Polymer nanofiber scaffolds and uses thereof

ABSTRACT

A polymer nanofiber scaffold includes a plurality of melt extruded nanofibers that are chemically modified to append surface functionality to the nanofibers.

RELATED APPLICATION

This application claims priority to U.S. Provisional Ser. No.62/489,263, filed Apr. 24, 2017, this application is also aContinuation-in-Part of U.S. Ser. No. 15/118,030, filed Aug. 10, 2016,which is a National Phase Filing of PCT/US2015/015243, filed Feb. 10,2015, which claims priority to U.S. Provisional Application Ser. No.61/937,756, filed Feb. 10, 2014, the subject matter of which isincorporated herein by reference in its entirety.

GOVERNMENT FUNDING

This invention was made with government support under Grant No.R00EB011530 awarded by The National Institutes of Health and DMR 0423914awarded by The National Science Foundation. The United States governmenthas certain rights to the invention.

BACKGROUND

Traditional industrial processes for synthetic polymer microfiberspinning can be classified as either solvent-based or melt-based. As thename implies, solvent processing involves the spinning of a polymersolution with solidification of the fiber either through coagulation ina non-solvent (wet-spinning) or solvent evaporation (dry-spinning). Incontrast, melt spinning produces fibers via the spinning of moltenpolymer that solidifies upon cooling; drawing usually accompanies thismelt-based process to induce chain orientation and enhance mechanicalproperties. Typically, accessing nanoscale cross-sections is difficult,where fiber diameters of only 10-20 μm are achieved with applications inthe textile industry. Pushing the limits of fiber production to thenanoscale has garnered recent attention in the processing arena.

Electrospinning is perhaps the most well-known, and one of the oldesttechniques for generating sub-micron fibers in lab-scale from a polymersolution, or less commonly, a polymer melt via the application of alarge electric field. This charged polymer jet is subjected toelectrostatic forces, which act to elongate, thin, and solidify thepolymer fiber in the characteristic “whipping instability” region.Although some success has been achieved with electrospun fibers inhigh-value added applications, such as air filtration, topical drugdelivery, and tissue engineering scaffolds, significant disadvantagesare low throughput and scalability. Additionally, electrospinningnecessitates large volumes of toxic solvents that must be recovered byspecialized equipment to make the process viable on a large scale.

Other approaches to nanofiber fabrication have emerged, includingrotary-jet spinning, gas jet blowing, melt blowing, and bicomponentfiber spinning. Recent advances in rotary jet spinning have focused on amelt-based approach to nanofiber production with throughputssignificantly higher than electrospinning, but improvements are ongoingto address processing complexity as it relates to broad applicability toa range of polymer systems. Melt blowing is a particularly commerciallyrelevant and scalable technique for achieving fiber diameters on theorder of tens of microns and higher; in this process, fibers aregenerated in-line by extrusion of a polymer through a die orifice, whilea hot air jet blows down the extrudate. It is process compatible with awide range of polymers, and is a solvent-less andenvironmentally-friendly manufacturing method. However, the pursuit ofnanoscale fibers has been limited primarily to polypropylene for airfiltration. Collectively, these limitations on nanofiber scalabilityincrease manufacturing costs and lower productivity.

Polymeric materials have become ubiquitous in regenerative medicine asscaffolds for cell-seeding, where they have found application in theinduction of cellular adhesion, proliferation, and differentiation.Nano-fibrous scaffolds are of particular use as they are porous,allowing transport of nutrients and waste products, have high surfacearea to volume ratios, and can provide directed cell growth based onfiber alignment. Synthetic fibers for regenerative medicine are usuallycomprised of polyesters, often poly(lactic acid) (PLA),poly(lactic-co-glycolic acid) (PLGA) or poly(caprolactone) (PCL), due totheir degradability via hydrolytic pathways and resultant non-toxicbyproducts. However, most polymeric scaffolds are unable to promotebiological effects, as synthetic polymers do not possess the biochemicalcues that are necessary to impact a cell's fate.

Modification of these polyester fibers typically relies on thedegradation of the polymer chains, either through hydrolysis to exposecarboxylic acids and alcohols, or through aminolysis to expose a secondfunctional group off of the amine. Both of these routes degrade thepolymer, potentially resulting in reduced mechanical properties andincreased erosion of the fibers. Recent work has aimed to amelioratethis through the synthesis of telechelic polymers, which could then beprocessed into a scaffold.

SUMMARY

Embodiments described herein relate to a polymer nanofiber scaffolds andto their use in, for example, tissue engineering, drug delivery, woundhealing, chemical processing, nanoelectronics, fabrics, and filtrationapplications. In some embodiments, a polymer nanofiber scaffold caninclude a plurality of melt extruded polymer nanofibers. The nanofiberscan each have a rectangular cross-section defined in part by anencapsulating polymer material that is separated from the nanofibers.The nanofibers include a plurality of click-reactive functional groupsof specific binding pairs extending from portions of outer surfaces ofthe nanofibers. The functional groups can be chemically bound to thenanofibers without degrading polymer chains of the nanofibers. Thefunctional groups can be appended to complementary click-reactive groupsof the specific binding pair that are conjugated to at least one agent.

In some embodiments, the concentration of functional groups extendingfrom the at least one portion can be at least about 0.1 nmol/cm². Theagent conjugated to complementary click-reactive group, which can beappended to the functional group, can include at least one of abioactive agent, diagnostic agent, therapeutic agent, catalyst, chargedmolecule, peptide, polypeptide, nucleic acid, polynucleotide, smallmolecule, nanoparticle, antibody, carbohydrate, or vector.

In other embodiments, the functional groups can be spatially arranged onthe nanofibers such that a first portion of the nanofibers has a firstconcentration of functional groups and a second portion of thenanofibers has a second concentration of functional groups differentthan the concentration of the first portion, e.g., different portions ofthe nanofibers can have different concentrations of the same ordifferent functional groups. In some embodiments, the functional groupscan be arranged on the nanofibers in a concentration gradient.

In still other embodiments, the plurality of click-reactive functiongroups can include first click reactive functional groups and secondclick reactive functional groups different than the first click reactivefunctional groups.

In some embodiments, the functional groups are chemically bound to thenanofibers with diarylhydroxymethylene linkages that are formed byreaction of click-reactive functional group substituted diarylketoneswith the polymer chains of the nanofibers. The click-reactive functionalgroups can include at least one of an amine, sulfate, thiol, hydroxyl,azide, alkyne, alkene, carboxyl groups, aldehyde groups, sulfone groups,vinylsulfone groups, isocyanate groups, acid anhydride groups, epoxidegroups, aziridine groups, episulfide groups, —CO₂N(COCH₂)₂,—CO₂N(COCH₂)₂, —CO₂H, —CHO, —CHOCH₂, —N═C═O, —SO₂CH═CH₂, —N(COCH)₂,—S—S—(C₅H₄N) and groups of the following structures:

wherein R₁ is hydrogen or C₁ to C₄ alkyl.

In other embodiments, the nanofibers can be formed of a biocompatible orcytocompatible polymer, such as a polyester (e.g., polycaprolactone).

Other embodiments described herein relate to a method of forming anagent functionalized polymer nanofiber scaffold. The method includesproviding a plurality of melt extruded polymer nanofibers. Thenanofibers can each have a rectangular cross-section defined in part byan encapsulating polymer material that is separated from the nanofibers.A plurality of click-reactive functional groups of a specific bindingpair can then be chemically bound to the fibers without degradingpolymer chains of the nanofibers. The click-reactive functional groupscan extend from portions of outer surfaces of the nanofibers. At leastone agent is appended to the nanofibers by reacting the agent conjugatedto a complementary click-reactive group of the specific binding pairwith the click-reactive functional groups of the nanofibers.

In some embodiments, the agent can include at least one bioactive agent,diagnostic agent, therapeutic agent, catalyst, charged molecule,peptide, polypeptide, nucleic acid, polynucleotide, small molecule,nanoparticle, antibody, carbohydrate, or vector.

In other embodiments, the nanofibers can be formed by coextruding afirst polymer material with a second polymer material to form acoextruded polymer film having discrete overlapping layers of polymericmaterial; multiplying the overlapping layers to form a multilayeredcomposite film; and separating the first polymer material from thesecond polymer material to form the plurality of nanofibers having therectangular cross-section. Separating the polymer materials can include,for example, subjecting the multilayered composite film to a highpressure water stream or a high pressure air stream, or dissolving thesecond polymer material.

In some embodiments, the functional groups are spatially arranged on thenanofibers such that a first portion of the nanofibers has a firstconcentration of functional groups and a second portion of thenanofibers has a second concentration of functional groups differentthan the concentration of first portion. The functional groups can beappended to complementary click-reactive groups that are conjugated toat least one agent to provide a first concentration of agents on thefirst portion and a second concentration of agents on the secondportion.

In other embodiments, the functional groups can be spatially arranged onthe nanofibers such that different portions of the nanofibers havedifferent concentrations of the functional groups. The functional groupscan be appended to complementary click-reactive groups that areconjugated to at least one agent to provide different concentrations ofagents on different portions of the nanofibers.

In still other embodiments, the plurality of click-reactive functiongroups can include first click reactive functional groups and secondclick reactive functional groups different than the first click reactivefunctional groups. The first click reactive functional groups canappended to first agents and the second click reactive functional groupscan be appended to second agents different than the first agents.

In some embodiments, the concentration of the functional click-reactivefunction groups and appended agent extending from at least one portionof the nanofibers is at least about 0.1 nmol/cm². The appended agent canbe a bioactive agent that promotes at least one of cell adhesion,growth, or proliferation and the polymer scaffold can be used in atleast one of biomedical, tissue engineering, and/or wound healingapplications, such as a wound dressing or engineered tissue construct.

BRIEF DESCRIPTION OF THE DRAWINGS

FIG. 1 is a schematic illustration of a coextrusion and layermultiplying process used to form a multilayered polymer composite filmin accordance with an embodiment.

FIG. 2 is a schematic illustration of coextruding skin layers onto thecomposite film of FIG. 1 to form a composite stream.

FIG. 3 is a schematic illustration of additional layer multiplying stepsfor the composite stream of FIG. 2.

FIG. 4A is a schematic illustration of stretching the composite streamof FIG. 2.

FIG. 4B is a schematic illustration of compressing the composite streamof FIG. 2.

FIG. 4C is a schematic illustration of delaminating the composite streamof FIG. 2.

FIG. 5 is a schematic illustration of delaminating a composite stream toform a scaffold.

FIG. 6 is a schematic illustration of a nanofiber surface functionalizedwith two different chemistries

FIGS. 7(A-B) illustrate: (A) schematic of co-extrusion and 2-dimensionalmultiplication system for producing nanofibers (top); and (B) scanningelectron micrograph of the as-extruded nanofibers following PEOdissolution (bottom). Scale bar (left)=20 μm, right=50 μm.

FIG. 8 illustrates a chemical scheme for modification of PCL nanofibers.

FIGS. 9(A-C) illustrate fluorescent confocal micrographs of PCLnanofibers. A) PCL nanofiber control with no CuSO₄ added during reactionwith AF₄₈₈. Scale bar=50 μm. B) PCL after the CuAAC reaction with AF₄₈₈,including CuSO₄. Scale bar=50 μm. C) Fluorescent intensity in the regionof interest, as indicated by the red lines in images A and B. The redline on the graph corresponds to image A, and the black line isindicative of image B.

FIGS. 10(A-F) illustrate: A) Full XPS spectrum of PCL, PCL-PrBz, andPCL-RGD. B) N_(1s) XPS spectrum of PCL nanofibers and PCL-RGD fibers, C)Confocal fluorescent microscopy image of NIH 3T3 cells after 72 hours ofgrowth on PCL-RGD scaffold. 10× objective. D) Confocal fluorescentmicroscopy image of NIH 3T3 cells after 72 hours of growth on PCL-RGDscaffold. 40× objective. E) Confocal fluorescent microscopy image of NIH3T3 cells after 72 hours of growth on control PCL scaffold. 10×objective. F) Confocal fluorescent microscopy image of NIH 3T3 cellsafter 72 hours of growth on control PCL scaffold. 40× objective. Blueindicates DAPI stain and green indicates actin green stain in confocalmicrographs.

FIG. 11 illustrates a schematic of a synthesis scheme to generate a PCLfiber gradient.

FIGS. 12(A-C) illustrate: A) schematic of gradient photomask as inkjetprinted onto transparency sheets: (a) is 100% black, (b) is 50% of blackand (c) is transparent as indicated in the text; B) Fluorescence imageof PCL-AF488 fiber gradient; and C) plot of fluorescence intensity overthe total fiber distance, indicating an approximate linear gradient ofintensity.

FIGS. 13(A-C) illustrate: A) Digital image of PCL gradient-modifiedIKVAV fibers, indicating points where IR spectra were taken. B) ATR-FTIRspectrum of varying spots on the PCL gradient (indicated in A). Thearrow indicates the amide I region derived from the IKVAV peptide. C)ATR-FTIR imaging at 1628 cm⁻¹ (scale bar=50 μm, intensity is indicatedby a heat map as is indicated on the right).

FIGS. 14(A-C) illustrate: A) Digital image of IKVAV gradient fibers,indicating spots taken for water contact angle and X-ray photoelectronspectroscopy. B) Water contact angle)(°) of PCL-g-IKVAV fibers with fourdistinct spots from left (low peptide concentration) to right (highpeptide concentration). Error bars represent standard deviation (n=3).C) Full XPS wide scan from left to right of PCL-gradient (spots 1-4), asindicated in A. IKVAV fibers shows increasing intensity of nitrogen(N1s) with increasing UV irradiation intensity.

FIGS. 15(A-I) illustrate confocal microscopy images of PC-12 cells.Cells were cultured for 5 days and stained with DAPI and antiβ-III-tubulin. Fiber orientation is approximately horizontal. A-C) PC-12cells cultured on unmodified PCL, corresponding to spots (a), (b) and(c). D-F) PC-12 cells cultured on PCL-ng-IKVAV which correspond spot(a), (b) and (c) in FIG. 12A. G-I) PC-12 cells cultured on 3 differentregions of PCL-g-IKVAV which correspond spot (a), (b) and (c). Thedirection of the gradient is left to right, where G is the lowestpeptide concentration and I is the highest. (Scale bar=50 μm).

FIGS. 16(A-C) illustrate non-gradient photomasks which correspond todifferent intensities of the gradient photomask. A) 100% black photomaskcorresponding to the least amount of UV fluence in the gradientphotomask. B) 50% black photomask where UV intensity corresponds to theexact center of the gradient photomask. C) Transparent photomaskcorresponding to the highest UV fluence through the gradient photomask.

FIGS. 17(A-C) illustrate images showing AF₄₈₈ clicked PCL fiberbundles;Top: Digital image of fibers corresponding to the individual photomasksin FIG. 16. Groups A, B, and C fibers used photomask A, B, and C toperform photochemistry with Pr-Bz before click chemistry. The tableindicates the intensity of UV light and the surface coverage of dyesconjugated onto fibers (n=3).

FIGS. 18(A-D) illustrate (A) EGF-MMP, (B) EGF control protein, (C)SDSPAGE gel of purified control EGF (7.34 kDa) and EGF-MMP (8.16 kDa),(D) MALDI-TOF MS analysis of control EGF protein and EGF-MMP.

FIGS. 19(A-B) illustrate (A) MTT cell viability assay of HaCaT (humankeratinocytes) and A431 (epidermoid carcinoma) cells in the presence ofEGF-MMP (B) MALDI-TOF MS indicating the molecular weight of EGF-MMPprotein before and after MMP-9 cleavage.

FIGS. 20(A-D) illustrate (A) SEM micrograph, fiber thickness=2.6±1.5 μm,Scale Bar=10 μm. (B) Pore size distribution of the fiber mat as measuredvia porimeter, mean pore diameter=25.6 μm. (C) ATR-FTIR spectrum ofEGF-MMP conjugated to fiber mats. Black=unirradiated, Red=irradiated.(D) Kinetic study of released EGF and EGF-MMP from fiber mats afterMMP-9 cleavage.

FIGS. 21(A-B) illustrate (A) Optical micrographs of gap distance betweenHaCaT cells at 0 and 24 hours after scratch test (scale bar=200 μm). (B)Relative scratch closure distance after 24 hours. Error bars areexpressed as standard deviations.

FIGS. 22(A-B) illustrate proposed mechanism for site-specificmodification of the protein in the presence of PLP. MALDI-TOF massspectrum indicates protein (A) at 8162 m/z and (B) at 8409 m/z, matchingmodel molecular weights, indicating modified EGF-MMP.

FIGS. 23(A-B) illustrate (A) Scheme of oxime ligation of the TAMRA dyewith the ketone-modified EGF protein, (B) FPLC trace of control EGF andEGF-MMP (left); FPLC trace of ketone modified control EGF and EGFMMP(middle); FPLC trace of TAMRA conjugated control EGF and EGF-MMP(right).The maximum absorbance of aminooxyl-5(6)-TAMRA is shown at 555 nm (red).The other wavelengths monitored were 280 nm (black) and 260 nm (blue).

FIG. 24 illustrates ATR-FTIR spectra of EGF-MMP conjugation to PCL fibermat. The spectrum from the back of the fiber mat is shown in black (noUV irradiation) and the spectrum from the front is shown in red (UVirradiation).

FIG. 25 illustrates bioconjugation of EGF to an aminooxy decorated PCLfiber mat and MMP-mediated release.

DETAILED DESCRIPTION

Embodiments described herein relate to surface functionalized polymernano-fibrous (or nanofiber) scaffolds, methods of forming the surfacefunctionalized polymer nanofiber scaffolds, and to the use of thescaffolds in, for example, tissue engineering, drug delivery, woundhealing, chemical processing, nanoelectronics, fabrics, and filtrationapplications. The nanofiber scaffolds can include a plurality ofnon-woven or woven nanofibers that are formed from commodity polymersusing a continuous extrusion process. The nanofibers can have arectangular cross-section of about 10 nm (height)×10 nm (width) to about10 μm×10 μm, with variations in between, and a surface area of at leastabout 1 cm²/g or more. The nanofibers and/or scaffold formed from thenanofibers can be chemically modified (or surface modified)non-destructively after formation to chemically bond click reactivefunctional groups of specific binding pairs onto the nanofibers suchthat the click-reactive functional groups extend uniformly across thesurface of the nanofibers and/or scaffold or from selected portions ofthe nanofibers or scaffold at varying concentrations, types, and/ordensities. Complementary click-reactive groups of the specific bindingpairs conjugated to any number of any number of agents and/or chemicalentities can be click-reacted with the click-reactive groups chemicallybound to the nanofibers to the agents to the nanofibers. The agentsappended to the click reactive groups of the fibers can include, forexample, bioactive agents, diagnostic agents, therapeutic agents,catalysts, charged molecules, peptides, polypeptides, nucleic acids,polynucleotides, small molecules, nanoparticles, antibodys,carbohydrates, and vectors.

In some embodiments, one or more similar or different functional groupscan be spatially arranged on the nanofibers such that different portionsof the nanofibers have different concentrations of the functionalgroups. For example, a plurality of first functional groups can bearranged on the nanofibers in a first concentration gradient and/or aplurality of second functional groups can be arranged on the nanofibersin a second concentration gradient.

Advantageously, the nanofiber scaffold can be formed or fabricated usingsolely commercially available polymers, such as PCL and poly (ethyleneoxide) (PEO). The process used to form the nanofiber scaffold can besolvent-free, allow for controllable cross-sectional dimensions of thefibers, and use FDA-friendly polymers during processing. The fabricationprocess is flexible because it involves the use of an extrusion linethat is composed of several basic multipliers. Arrangement of thesemultipliers allows control over the number, as well as the dimensions,of fibers contained in one extrudate. In addition, the cross-sectionalgeometry of the nanofibers is rectangular creating greater surface areato volume ratios (e.g., at least about 1 cm²/mg, at least about 10cm²/mg, at least about 20 cm²/mg, at least about 40 cm²/mg, at leastabout 50 cm²/mg or more), when compared to cylindrical fibers. Theincreased surface area can allow for a higher concentration of surfacemodifications to be available on the fiber, potentially improving thedisplay of chemical or biochemical cues.

The surface functionalized polymer nanofiber scaffold can be formed froma multilayered polymer composite film. FIG. 1 illustrates a coextrusionand multilayering process used to form a multilayered polymer compositefilm 10. First, a first polymer layer 12 and a second polymer layer 14are provided. The first layer 12 is formed from a first polymericmaterial (a) and the second polymer layer 14 is formed from a secondpolymer material (b) that is substantially immiscible and has a similarviscosity with the first polymer material (a) when coextruded. It willbe appreciated that one or more additional layers formed from thepolymer materials (a) or (b) or a different polymer materials may beprovided to produce the multilayered polymer composite film 10.

The term “polymer” or “polymeric material” as used in the presentapplication denotes a material having a weight average molecular weight(Mw) of at least 5,000. Preferably the polymer is an organic polymericmaterial. The term “oligomer” or “oligomeric material” as used in thepresent application denotes a material with a weight average molecularweight of from 1,000 to less than 5,000. Such polymeric materials can beglassy, crystalline or elastomeric polymeric materials.

Examples of polymeric materials that can potentially be used for thefirst and second polymer materials (a), (b) include, but are not limitedto, polyesters such as polylactic acid, poly(lactic-co-glycolic acid),polycaprolactone (PCL); and poly(ethylene naphthalate) and isomersthereof, such as 2,6-, 1,4-, 1,5-, 2,7-, and 2,3-polyethylenenaphthalate; polyalkylene terephthalates such as polyethyleneterephthalate, polybutylene terephthalate, andpoly-1,4-cyclohexanedimeth-ylene terephthalate; polyimides, such aspolyacrylic imides; polyetherimides; styrenic polymers, such as atactic,isotactic and syndiotactic polystyrene, α-methyl-polystyrene,para-methyl-polystyrene; polycarbonates, such asbisphenol-A-polycarbonate (PC); polyethylenes, such as polyethyele oxide(PEO); poly(meth)acrylates, such as poly(isobutyl methacrylate),poly(propyl methacrylate), poly(ethyl methacrylate), poly(methylmethacrylate), poly(butyl acrylate) and poly(methyl acrylate) (the term“(meth)acrylate” is used herein to denote acrylate or methacrylate);cellulose derivatives, such as ethyl cellulose, cellulose acetate,cellulose propionate, cellulose acetate butyrate, and cellulose nitrate;polyalkylene polymers, such as polyethylene, polypropylene,polybutylene, polyisobutylene, and poly(4-methyl)pentene; fluorinatedpolymers such as perfluoroalkoxy resins, polytetrafluoroethylene,fluorinated ethylene-propylene copolymers, polyvinylidene fluoride, andpolychlorotrifluoroethylene and copolymers thereof; chlorinatedpolymers, such as polydichlorostyrene, polyvinylidene chloride andpolyvinylchloride; polysulfones; polyethersulfones; polyacrylonitrile;polyamides such as nylon, nylon 6,6, polycaprolactam, and polyamide 6(PA6); polyvinylacetate; and polyether-amides. Also suitable arecopolymers, such as styrene-acrylonitrile copolymer (SAN), preferablycontaining between 10 and 50 wt %, preferably between 20 and 40 wt %,acrylonitrile, styrene-ethylene copolymer; andpoly(ethylene-1,4-cyclohex-ylenedimethylene terephthalate) (PETG).Additional polymeric materials include an acrylic rubber; isoprene (IR);isobutylene-isoprene (IIR); butadiene rubber (BR);butadiene-styrene-vinyl pyridine (PSBR); butyl rubber; chloroprene (CR);epichlorohydrin rubber; ethylene-propylene (EPM);ethylene-propylene-diene (EPDM); nitrile-butadiene (NBR); polyisoprene;silicon rubber; styrene-butadiene (SBR); and urethane rubber. Additionalpolymeric materials include block or graft copolymers. In one instance,the polymeric materials used to form the layers 12, 14 may constitutesubstantially immiscible thermoplastics. In addition, each individuallayer 12, 14 may include blends of two or more of the above-describedpolymers or copolymers, preferably the components of the blend aresubstantially miscible with one another yet still maintainingsubstantial immiscibility between the layers 12, 14. The componentscomprising the layers 12, 14 can include organic or inorganic materials,including nanoparticulate materials, designed, for example, to modifythe mechanical properties of the components, e.g., tensile strength. Itwill be appreciated that potentially any extrudable polymer can be usedas the first polymer material (a) and the second polymer material (b) solong as upon coextrusion such polymer materials (a), (b) aresubstantially immiscible and form discrete layers or polymer regions.Such materials can have a substantially similar viscosity uponcoextrusion.

In some embodiments, the polymers used to for the first and/or secondpolymer can be biodegradable and/or substantially biocompatible orcytocompatible (i.e., substantially non-cytotoxic). The use ofbiodegradable and substantially biocompatible or cytocompatible polymersallows a surface functionalized nanofiber scaffold to be formed that canbe used in medical applications, e.g., tissue engineering or woundhealing. Examples of that polymers that are substantially biocompatibleor cytocompatible include polyesters, such as PCL, paired with PEO.

Referring to FIG. 1, the layers 12, 14 are co-extruded and multiplied inorder to form the multilayered polymer composite film 10. In particular,a pair of dies 40, 50 is used to coextrude and multiply the layers 12,14. Each layer 12, 14 initially extends in the y-direction of an x-y-zcoordinate system. The y-direction defines the length of the layers 12,14 and extends in the general direction of flow of material through thedies 40, 50. The x-direction extends transverse, e.g., perpendicular, tothe y-direction and defines the width of the layers 12, 14. Thez-direction extends transverse, e.g., perpendicular, to both thex-direction and the y-direction and defines the height or thickness ofthe layers 12, 14.

The layers 12, 14 are initially stacked in the z-direction and define aninterface 20 therebetween that resides in the x-y plane. As the layers12, 14 approach the first die 40 they are separated from one anotheralong the z-axis to define a space 22 therebetween. The layers 12, 14are then re-oriented as they pass through the first die 40. Morespecifically, the first die 40 varies the aspect ratio of each layer 12,14 such that the layers 12, 14 extend longitudinally in the z-direction.The layers 12, 14 are also brought closer to one another until theyengage or abut one another along an interface 24 that resides in the y-zplane.

The layers 12, 14 then enter the second die 50 where layermultiplication occurs. The second die 50 may constitute a single die orseveral dies which process the layers 12, 14 in succession (not shown).Each layer 12, 14 is multiplied in the second die 50 to produce aplurality of first layers 12 and a plurality of second layers 14 thatalternate with one another to form the multilayered polymer compositefilm 10. Each pair of layers 12, 14 includes the interface 24 thatresides in the y-z plane. The layers 12, 14 are connected to one anothergenerally along the x-axis to form a series of discrete, alternatinglayers 12, 14 of polymer material (a), (b). Although three of each layer12 and 14 are illustrated it will be appreciated that the multilayeredpolymer composite film 10 may include, for example, up to thousands ofeach layer 12, 14.

Referring to FIG. 2, once the multilayered polymer composite film 10 isformed a detachable skin or surface layer 30 is applied to the top andbottom of the film 10 such that the film 10. In particular, themultilayered polymer composite film 10 enters a die 60 where the film 10is sandwiched between two skin layers 30 along the z-axis to form afirst composite stream 100. The skin layer 30 may be formed from thefirst polymer material (a), the second polymer material (b) or a thirdpolymer material (c) different from the first and second polymermaterials (a), (b). One or both of the skin layers 30 may, however, beomitted (not shown).

Referring to FIG. 3, the first composite stream 100 is divided along thex-axis into a plurality of branch streams 100 a, 100 b and processedthrough a pair of multiplying dies 70, 80. In the die 70, the streams100 a, 100 b are stacked in the z-direction, stretched in both thex-direction and the y-direction, and recombined to form a secondcomposite stream 110 that includes a plurality of multilayered films 10alternating with skin layers 30. Biaxial stretching of the branchstreams 100 a, 100 b in the x-direction and y-direction may be symmetricor asymmetric.

The die 80 performs similar modifications to the second composite stream110 that the die 70 performed on the branch streams 100 a, 100 b. Inparticular, in the die 80 the second composite stream 110 is dividedalong the x-axis, stacked along the z-axis, stretched in both thex-direction and the y-direction, and stacked in the z-direction to forma third composite stream 120. The third composite stream 120 shown inFIG. 3 includes four multilayered composite films 10 that alternate withfive skin layers 30, although more or fewer of the films 10 and/orlayers 30 may be present in the third composite stream 120. Regardless,the third composite stream 120 includes a plurality of layer interfaces24 between the layers 12, 14.

By changing the volumetric flow rate of the polymer layers 12, 14through the dies 70, 80, the thickness of both the polymer layers 12, 14and each multilayered polymer film 10 in the z-direction can beprecisely controlled. Additionally, by using detachable skin layers 30and multiplying the composite streams 100, 110 within the dies 70, 80,the number and dimensions of the layers 12, 14, the multilayered polymerfilm 10, and the branch streams 100 a, 100 b in the x, y, andz-directions can be controlled.

Referring to FIGS. 4A and 4B, the first composite structure 100 may bemechanically processed by, for example, at least one of orientation(FIG. 4A), compression (FIG. 4B), and ball-mill grinding (not shown). Asshown, the composite stream 100 is stretched in the y-direction asindicated generally by the arrow “S”, although the composite stream 100may alternatively be stretched in the x-direction (not shown). FIG. 4Billustrates the composite stream 100 being compressed in the z-directionas indicated generally by the arrow “C”. The degree of stretching and/orcompression will depend on the application in which the nanofibersand/or scaffold formed from the multilayered polymer film 10 is to beused. The ratio of y-directional stretching to z-direction compressionmay be inversely proportional or disproportional.

In one embodiment, the multilayer film can be uniaxially stretched inthe y-direction or S-direction at a draw ratio of about 1 to about 10 todecrease the cross-sectional dimension of the fibers and increase thesurface area of the nanofibers and scaffold so formed.

Referring to FIG. 4C, the first composite stream 100 can be furtherprocessed to cause the components 12, 14, 30 thereof to separate ordelaminate from one another and form a plurality of fibers or fiber-likestructures 12 a, 14 a from the layers 12, 14. The removed skin layers 30are discarded. In one instance, the layers 12, 14, 30 are mechanicallyseparated by high pressure water jets (not shown). In particular, twoopposing ends of the composite stream 100 can be fixed and water jetswith a nozzle pressure of no less than about 2000 psi can be applied tothe composite stream 100 to separate the layers 12, 14, 30 completely,thereby forming the nano-fibers 12 a, 14 a. More specifically, applyinghigh pressure water to the first composite stream 100 removes theinterfaces 24 between the layers 12, 14, i.e., delaminates themultilayered polymer composite films 10, to form the fibers 12 a and 14a. Although delamination of the first composite stream 100 isillustrated, it will be appreciated that the multilayered polymercomposite film 10, the second composite stream 110 or the thirdcomposite stream 120 may likewise be delaminated via high pressure wateror the like to form the fibers 12 a, 14 a.

Alternatively, the polymer material (a) or (b) of one of the layers 12,14 is selected to be soluble in a particular solvent while the otherpolymer material (a) or (b) is selected to be insoluble in that solvent.Accordingly, immersing the composite stream 100 in the solvent separatesthe layers 12, 14 by wholly removing, e.g., dissolving, not only theinterfaces 24 between the layers 12, 14 but removed the soluble layers12 or 14 entirely. The insoluble layers 12 or 14 are therefore leftbehind following solvent immersion. The same solvent or a differentsolvent may be used to dissolve the skin layers 30, when present. Theremaining soluble layers 12 or 14 form the fibers 12 a or 14 a. In oneinstance, the solvent can be water and in some instances no organicsolvent is used.

Optionally, as illustrated in FIG. 5, a multilayer composite stream thatincludes a polymer material soluble in a particular solvent and anotherpolymer material insoluble in that solvent can be bundled orconsolidated prior to solvent immersion. Immersing the bundled compositestream in the solvent separates the layers and forms a scaffold of thepolymer nanofibers.

Whether the fibers 12 a and/or 14 a are formed by mechanicallyseparating the layers 12 or 14 or dissolving one of the layers 12 or 14with a solvent, the nanofibers 12 a and/or 14 a produced by thedescribed coextrusion process have rectangular cross-sections ratherthan the conventional, round cross-sections formed by electrospinning.These rectangular or ribbon-like nanofibers 12 a or 14 a have a largersurface area-to-volume ratio than round fibers developed using spinningmethods. Regardless of the method of separation enlisted, the nanofibers12 a and/or 14 a can stretch, oscillate, and separate from each other atthe interface 24. Furthermore, due to the aforementioned mechanicalprocessing techniques of FIGS. 4A and 4B, the exact cross-sectionaldimensions of the rectangular fibers 12 a and/or 14 a can be preciselycontrolled. For example, the rectangular fibers 12 a and/or 14 a can bemade smaller and strengthened via mechanical processing.

Due to the construction of the first composite stream 100 and the fixedsizes of the dies 40-80, the composition of the vertical layers 12, 14and surface layers 30 is proportional to the ratio of the height in thez-direction of a vertical layer 12, 14 section to that of a surfacelayer 30 section. Therefore, if the layer 12 (or 14) is selected to formthe rectangular fibers 12 a (or 14 a), the thickness and height of thefinal fibers 12 a (or 14 a) can be adjusted by changing the ratio of theamount of the layers 12, 14 as well as the amount of surface layer 30.For example, increasing the percentage of the amount of the material (b)of the layers 14 relative to the amount of the material (a) of thelayers 12 and/or increasing the amount of the material of the surfacelayers 30 results in smaller rectangular fibers 12 a. Alternatively, oneor more of the dies 40-80 may be altered to produce nanofibers 12, 12 a,14, 14 a having a size and rectangular cross-section commensurate withthe desired application. In one instance, one or more of the dies 40-80could be modified to have a slit or square die construction to embed thefibers 12, 12 a, 14, 14 a within the surface layers 30.

The extrusion process can be tailored to produce vertically layeredfilms 10 with designer layer/fiber thickness distributions. For example,the relative material compositions of the polymers (a), (b) of thelayers 12, 14 can be varied with great flexibility to producerectangular polymer fibers 12, 12 a, 14, 14 a with highly variableconstructions, e.g., 50/50, 30/80, 80/30, etc. The rectangular polymerfibers 12, 12 a, 14, 14 a can be highly oriented and strengthened bypost-extrusion orienting. Furthermore, a wide magnitude of layer 12, 14thicknesses in the z-direction is achievable from a few microns down totens of nanometers depending on the particular application.

The polymer nanofibers so formed can be consolidated to produce polymernanofiber scaffolds. In one embodiment, as shown in FIG. 5, thenanofibers can be consolidated by bundling the composite stream prior toseparation of the fibers or salvation of a polymer. In otherembodiments, the nanofibers can be consolidated by compressing, weaving,and physically mixing the fibers.

The nanofiber scaffolds can have various densities and porositiesdepending on the fiber cross-section and the process used to consolidatethe fibers. For example, the scaffolds can have a porosity of about 1%by volume to about 50% by volume, and a pore size of about 1 μm to about100 μm. In some embodiments, each fiber can have a rectangular crosssection of 10 nm (height)×10 nm (width) to about 10 μm×10 μm, withvariations in between. The nanofibers can have surface area of at leastabout 1 cm²/mg, at least about 10 cm²/mg, at least about 20 cm²/mg, atleast about 40 cm²/mg, at least about 50 cm²/mg or more.

The nanofibers of the scaffold can be chemically modified (or surfacemodified) non-destructively after processing to append click reactivefunctional groups onto the nanofibers such that the click-reactivefunctional groups extend from portions or selected portions of thenanofiber.

In some embodiments, the click-reactive functions groups can bechemically bound to the nanofibers with diarylhydroxymethylene linkagesthat are formed by reaction of click-reactive functional groupsubstituted diarylketones with polymer chains of the nanofibers. Forexample, modified diarylketones, such as benzophenones, which presentclick-reactive functional groups, such as alkynes, ketones, andalkoxyamines, can be attached to polymer chains of the nanofibers usingphotochemistry. In the presence of ultra-violet (UV) light, thediarylketones can generate radicals and undergo free radical insertionin the polymer backbone of the nanofibers.

For example, as shown below, photochemistry can used to chemically binda click-reactive group substituted benzophenone to PCL of a nanofiber:

wherein R is a click-reactive group.

By click-reactive functional group, it is meant that the functionalgroup has click reactivity and/or can undergo a click reaction with acomplementary click reactive functional group or molecule. Examples ofthe types of reactions that are known to have click reactivity includecycloaddition reactions. These reactions represent highly specificreactant pairs that have a chemoselective nature, meaning that theymainly react with each other and not other functional groups. Oneexample of a cycloaddition reaction is the Huisgen 1,3-dipolarcycloaddition of a dipolarophile with a 1,3 dipolar component thatproduce five membered (hetero)cycles. Examples of dipolarophiles arealkenes, alkynes, and molecules that possess related heteroatomfunctional groups, such as carbonyls and nitriles. Specifically, anotherexample is the 2+3 cycloaddition of alkyl azides and acetylenes. Othercycloaddition reactions include Diels-Alder reactions of a conjugateddiene and a dienophile (such as an alkyne or alkene).

Other examples of the types of reactions that are known to have clickreactivity include a hydrosilation reaction of H—Si and simplenon-activated vinyl compounds, urethane formation from alcohols andisocyanates, Menshutkin reactions of tertiary amines with alkyl iodidesor alkyl trifluoromethanesulfonates, Michael additions, e.g., the veryefficient maleimide-thiol reaction, atom transfer radical additionreactions between —SO₂Cl and an olefin, metathesis, Staudinger reactionof phosphines with alkyl azides, oxidative coupling of thiols, many ofthe procedures already used in dendrimer synthesis, especially in aconvergent approach, which require high selectivity and rates,nucleophilic substitution, especially of small strained rings like epoxyand aziridine compounds, carbonyl chemistry like formation of ureas, andaddition reactions to carbon-carbon double bonds like dihydroxylation.Therefore, the attached click-reactive functional group may be chosen,for example, from an alkyne group, acetylene group, an azido-group, anitrile group, acetylenic group, amino group, or phosphino group. Theclick chemistry reaction may results in the addition of a functionalgroup selected from amino, primary amino, hydroxyl, sulfonate,benzotriazole, bromide, chloride, chloroformate, trimethylsilane,phosphonium bromide or bio-responsive functional group.

In some embodiments, the click-reactive group can include at least oneof an amine, sulfate, thiol, hydroxyl, azide, alkyne, alkene, carboxylgroups, aldehyde groups, sulfone groups, vinylsulfone groups, isocyanategroups, acid anhydride groups, epoxide groups, aziridine groups,episulfide groups, —CO₂N(COCH₂)₂, —CO₂N(COCH₂)₂, —CO₂H, —CHO, —CHOCH₂,—N═C═O, —SO₂CH═CH₂, —N(COCH)₂, —S—S—(C₅H₄N) and groups of the followingstructures:

wherein R₁ is hydrogen or C₁ to C₄ alkyl.

The click-reactive functional group substituted diarylketones can beprovided on the nanofibers or nanofiber scaffold by coating a solutionof the click-reactive functional group substituted diarylketones ontothe nanofibers, removing excess solvent, and irradiating the coatednanofibers or scaffold with UV light. In some embodiments, the solutionof click-reactive functional group substituted diarylketones can becoated, e.g., spray coated, on the nanofibers of the scaffold such thatthe concentration of functional groups extending from at least oneportion is at least about 0.1 nmol/cm². The amount, concentration,and/or spatial location of the click-reactive functional groupsubstituted diarylketones provided on the nanofibers of the scaffold canbe controlled such that different densities or concentrations of theclick-reactive functional group substituted diarylketones are localizedor patterned on different portions of the scaffold. In some embodiments,surface coverage of the nanofibers can be, for example, about 0.1nmol/cm² to about 10 nmol/cm².

In some embodiments, one or more similar or different click-reactivefunctional groups can be spatially arranged on the nanofibers such thatdifferent portions of the nanofibers have different concentrations ofthe same or different click-reactive functional groups. For example, aplurality of first functional groups can be arranged on the nanofibersin a first concentration gradient and/or a plurality of secondfunctional groups can be arranged on the nanofibers in a secondconcentration gradient.

In some embodiments, gradient concentrations of the click-reactivefunctional groups can be provided on the fibers or scaffold by sprayingor coating different portions of the nanofibers and/or scaffolddifferent with different types or concentrations of click-reactivefunctional group substituted diarylketones. In other embodiments,gradient concentrations of the click-reactive functional groups can beprovided on the fibers or scaffold using a photomasking strategy topattern or modulate the concentration of click-reactive functional groupsubstituted diarylketones deposited on select portions of the fibers.For example, photomask having defined regions of transparency rangingfrom completely UV transparent to opaque can be used to pattern ordefine the surface location and concentration of the click-reactivefunctional group substituted diarylketones bound to the nanofibers ofthe scaffold.

The gradients can be comprised of different click-reactive functionalgroup substituted diarylketones bound to the nanofibers of the scaffold,such as benzophenones having different click-reactive functional groups(e.g., alkyne, aminoxy, etc.). As discussed below, for example, thedifferent click-reactive functional group substituted diarylketonesbound to the nanofibers can be provided in a particular pattern orconcentration on the nanofiber scaffold to allow the attachment ofdifferent agents to the nanofiber scaffold.

For example, as shown in FIG. 6, a first solution of ketoneclick-reactive group substituted diarylketones can be provided on a PCLnanofiber and irradiated with UV light through a photomask with definedtransparency regions to provide the PCL nanofiber with a definedconcentration gradient of the ketone click-reactive group substituteddiarylketones. A second solution of alkyne click-reactive groupsubstituted diarylketones can then be provided on the PCL nanofiber andirradiated with UV light through a photomask with defined transparencyregions to provide the PCL nanofiber with a defined concentrationgradient of the alkyne click-reactive group substituted diarylketones.

The click-reactive functional groups bound to the polymer chains of thenanofibers using diarylketone chemistry can participate inclick-reactions, such as cycloaddition or Michael addition reactions,with complementary click-reactive groups of specific binding pairs. Thecomplementary click-reactive groups of specific binding pairs can beconjugated to one more agents such that the agents can be readilyattached to the surface functionalized nanofiber using click-reactivechemistry. Examples of click-reactions used to conjugate agents includethe following:

wherein R₂ is one or more agent that can be conjugated to the surfacefunctionalized nanofiber.

Binding different click-reactive group substituted diarylketones to thenanofiber can allow the attachment of different agents conjugated todifferent complementary click-reactive groups. This allows one or moresimilar or different agents to be spatially arranged on the nanofiberssuch that different portions of the nanofibers have differentconcentrations of the click-reacted agent. For example, a plurality offirst agents can be arranged on the nanofibers in a first concentrationgradient and/or a plurality of agents can be arranged on the nanofibersin a second concentration gradient depending on the concentrationgradients of the bound click-reactive functional groups.

The agents attached to the surface functionalized nanofibers and/orscaffold using click-reactive chemistry can include any compound, smallmolecule, peptide, polypeptide, carbohydrate, nucleic acid, bioactiveagent, diagnostic agent, therapeutic agent, catalyst, charged molecule,nanoparticle, antibody, carbohydrate, cell, semiconductor, or vector orany other material that can modify at least one physical, chemical, orfunctional property of the nanofibers or scaffold. The specific agentbound to the surface functionalized nanofiber can depend on the specificapplication, use, or function of the nanofibers and/or nanofiberscaffold.

In some embodiments, the surface functionalized nano-fiber scaffolds canbe used in a variety of biomedical applications, including tissueengineering, drug delivery, wound healing, and regenerative medicineapplications. Advantageous, such scaffolds can have high surface area(e.g., greater than about 10 cm²/mg of fiber) and high aspect ratiofibers for promoting cell adherence, growth, proliferation, anddifferentiation as well as facilitate liquid and gas transport (e.g.,facilitate oxygenation of tissue and efflux of waste products from thetissue). The scaffolds can also include one or more bioactive agentsand/or therapeutic agents that provide improved and/or directed celladherence, growth, proliferation, and differentiation. Bioactive agentsthat can be click-reacted to the surface fuctionalized nanofiberscaffold can include any bioactive agent capable of promoting celladhesion, growth, proliferation, and differentiation, tissue formation,destruction, and/or targeting a specific disease state. Examples ofbioactive agents include chemotactic agents, various proteins (e.g.,short term peptides, bone morphogenic proteins, collagen, glycoproteins,and lipoprotein), cell attachment mediators, biologically activeligands, integrin binding sequence, various growth and/ordifferentiation agents and fragments thereof (e.g., EGF), HGF, VEGF,fibroblast growth factors (e.g., bFGF), PDGF, insulin-like growth factor(e.g., IGF-I, IGF-II) and transforming growth factors (e.g., TGF-βI-III), parathyroid hormone, parathyroid hormone related peptide, bonemorphogenic proteins (e.g., BMP-2, BMP-4, BMP-6, BMP-7, BMP-12, BMP-13,BMP-14), sonic hedgehog, growth differentiation factors (e.g., GDF5,GDF6, GDF8), recombinant human growth factors (e.g., MP-52 and the MP-52variant rhGDF-5), cartilage-derived morphogenic proteins (CDMP-1,CDMP-2, CDMP-3), small molecules that affect the upregulation ofspecific growth factors, tenascin-C, hyaluronic acid, chondroitinsulfate, fibronectin, decorin, thromboelastin, thrombin-derivedpeptides, heparin-binding domains, heparin, heparin sulfate,polynucleotides, DNA fragments, DNA plasmids, MMPs, TIMPs, interferingRNA molecules, such as siRNAs, DNA encoding for an shRNA of interest,oligonucleotides, proteoglycans, glycoproteins, and glycosaminoglycans.

In one example, the surface functionalized nanofiber scaffold can beused as a substrate for a wound dressing. The wound dressing can have alength, a width, and a thickness that may be varied depending upon theparticular application. For example, the wound healing dressing can havea sleeve-like or cylindrical configuration as well as a rectangular orsquare-shaped configuration. In one example, the wound healing dressingcan have a thickness of about 1 cm to about 10 cm or greater, a lengthof about 2 cm to about 30 cm or greater, and a width of about 2 cm toabout 30 cm or greater.

The wound dressing can include a plurality of peptides and proteinsclick-reacted to the surface of the scaffold and that can promote cellgrowth, attachment, and migration. An example of a protein that can beclick-reacted to the surface functionalized nanofiber scaffold isepidermal growth factor (EGF). EGF is known to enhance treatment ofchronic wounds. Examples of peptides that can be click-reacted to thesurface functionalized nanofiber scaffold include an RGD sequencederived from fibronectin, which is known to promote cellular adhesion,and an IKVAV peptide, which can be used to induce neural growth andmigration.

The EGF, RGD, and/or IKVAV can be provided or patterned in multiplegradients on the nanofiber scaffold to promote cell adherence, growth,migration. The gradients can run in the same or opposite directions. Forexample, the wound healing dressing formed from the surfacefunctionalized nanofiber scaffold can be can include gradients of theEGG, RGD, and/or IKVAV that facilitate cell growth, attachment,migration when the wound healing dressing is applied to tissue beingtreated in a subject in need thereof. In some embodiments a gradient ofIKVAV can be provided on the nanofibers and/or scaffold to promoteneural cell growth and migration.

It will be appreciated that the surface functionalized nanofibers and/orscaffold can be potentially used in any application where it isdesirable to modify at least one physical, chemical, or functionalproperty of the nanofibers and/or scaffold. For example, the surfacefunctionalized nanofibers can be used to form surface functionalizedmembrane supports in which various chemicals, catalysts, or other agentsare patterned on the surface of the membrane. Such membranes can beuseful in different processes, such as filtration, desalination, waterpurification, chemical processing or synthesis, nanoelectronics,non-woven fabrics, drug delivery, drug discovery, information storage,mechanical actuation, microarrays, and bioconjugation.

The following examples are included to demonstrate preferred embodimentsof the invention. It should be appreciated by those of skill in the artthat the techniques disclosed in the examples which follow representtechniques discovered by the inventor to function well in the practiceof the invention, and thus can be considered to constitute preferredmodes for its practice. However, those of skill in the art should, inlight of the present disclosure, appreciate that many changes can bemade in the specific embodiments which are disclosed and still obtain alike or similar result without departing from the spirit and scope ofthe invention.

Example 1

In this example, we used a melt co-extrusion in conjunction with amodular chemistry to yield polyester nanofibers with pendant surfacefunctionality. The processing method makes use of the co-extrusion ofPCL and PEO through a series of dye multipliers to form a nanofibrousPCL matrix within a PEO tape. PCL and PEO are melt-pumped and layered ontop of one another in the extrusion line (FIG. 7). From here a verticalmultiplier is used to rearrange the extrudate to yield a vertical layerstructure (Step A). This is then followed by a series of verticalmultipliers that cut the flow horizontally and expands the flow fieldsin the vertical direction to align in parallel. This process is repeatedeight times to yield a vertically aligned, layered flow comprised ofalternating PCL (512 layers) and PEO (512 layers) with 1024 total layers(Step B). The tape is then combined with two surface layers of PEO onthe top and bottom (Step C) and is split vertically with one sidestacked on top of the other (2×) to yield a PCL nanomatrix embedded in aPEO tape. This procedure is repeated one additional time, and anextrudate containing 512 PCL nanodomains embedded in a PEO matrix isobtained (Step D). The PEO in the tape can be removed by dissolution ina water bath or by using a high pressure water jet to produce a PCLnanofiber matrix. A scanning electron micrograph of the PCL fibers afterthe dissolving procedure shows fibers displaying cross-sectionaldimensions of approximately 400-1000 nm by 2-5 μm (FIG. 7). Afterremoval of the PEO matrix, <1% of PEO remained as determined by NMR andDSC (Supporting Information), yielding a scalable PCL nanofiber mat.

Once the fibers were extruded and separated, we sought to further modifythe fiber mats to add pendant functionality. A number of papers havefocused on the aminolysis of polyester fibers, in which a bifunctionalamine is used to introduce a reactive group for further derivatization.In the case of the extruded nanofibers, prepared as described, theaminolysis reaction using either propargylamine or hexamethylene diamineshowed minimal surface modification. This could be due to the smallfraction of PEO remaining on the exterior of the fibers, occludingaccess to the aminolysis pathway. Rather than pursuing forcing reactioncondition, an alternate route was chosen so as not to degrade thepolymer chains within the fibers. As such, a photochemical radicalreaction was employed to perform a C—H bond insertion into the backboneof the PCL polymer chains (FIG. 8). Benzophenone is commonly used toperform photochemical C—H insertions and has been previously used tomodify PCL surfaces. Propargyl benzophenone was prepared in high-yieldby reacting 4-hydroxyl benzophenone with propargyl bromide under basicconditions. The propargylbenzophenone derivative was solvent cast ontothe PCL fibers, irradiated via UV (33.2 mW/cm², 30 min on each side,320-500 nm) and washed with methanol to yield alkyne-decorated PCLnanofibers. The propargyl group could then be used in the CuAAC reactionto decorate the fiber with any azide containing molecule.

To probe the CuAAC chemistry, we first modified the PCL-alkyne fiberswith an azide containing fluorescent dye, AzideFluor488 (AF₄₈₈). Thefibers were modified using aqueous conditions optimized for theligand-accelerated CuAAC reaction, usingtris(3-hydroxypropyltriazolylmethyl)amine (THPTA) as the ligand toaccelerate the reaction. After the CuAAC reaction, upon visualinspection, the fibers were noticeably red after using standard reactionconditions. While in the absence of copper, there was no visible changethat could be attributed to a hydrophobic physical adsorption of the dyeonto the fiber. To further probe the surface coverage of the PCL fibers,confocal fluorescent microscopy was used to qualitatively andquantitatively evaluate the surface coverage of the fibers. As seen inthe case with no copper catalyst, negligible fluorescence is visible inthe micrograph (FIG. 9A), indicating that little dye non-specificallyadsorbs to the fiber bundle. However, in the presence of copper asignificant fluorescent intensity appears in the micrograph indicatingthat the chemistry is specific, rather than simple adsorption. Whenfluorescence intensity was quantified, modified fibers showedapproximately two-orders of magnitude higher fluorescent intensityrelative to control fibers (FIG. 9C).

In order to effect biological outcomes, it is necessary that asignificant surface coverage of the fibers be attained. To quantifythis, we determined both the total dye loading of the fibers inconjunction with surface area measurements. Brunauer Emmett Teller (BET)measurements were used to determine the surface area of the extrudedfibers, and UV-visible measurements were used to quantify dye loading.The BET technique relies on the surface adsorption of gas molecules; thetotal mass of the gas adsorbed allows one to calculate the area of thesurface. BET measurements were carried out, showing the surface are ofthe fibers to be 41.2 cm²/mg of fiber. Of note, electrospun fibers ofsimilar cross sectional dimensions showed an approximately 20-fold lowersurface area. This can be attributed to the rectangular geometry of theextruded fiber as compared to the electrospun cylindrical fiber. Oncethe surface area was determined, we labeled the fibers with AF₄₈₈, asdescribed. The fibers were dissolved in DCM and the absorbance at 501 nmwas compared to a standard curve of AF₄₈₈ to determine total surfaceloading of the fibers. The PCL-AF488 fibers were decorated withapproximately 15.5 nmol/mg of fiber, resulting in a surface coverage of0.38 nmol/cm². The surface coverage of the fibers is of particularimportance as a scaffold for regenerative medicine. A common peptide,and one we employed here, is the RGD motif. RGD sequences bind tointegrin receptors, and when immobilized on a surface, can lead to celladhesion and spreading, a key feature for tissue engineering scaffolds.Previous work showed that low pM/cm² surface coverages of RGD peptideson polymeric surfaces were sufficient to promote cell adhesion. Ourconcentrations are three-orders of magnitude higher than those necessaryfor adhesion, representing a viable scaffold to present biochemical cuesand for cell-seeding.

Finally, we sought to adhere an RGD peptide to promote cellular adhesiononto the PCL fiber scaffold. We first synthesized an RGD peptide with anN-terminal azide group (N₃-RGD), for attachment to the propargyl PCLfiber. The RGD peptide motif was chosen for its ability to promoteadhesion through the interaction with integrin cell surface receptors,as is known to occur with NIH 3T3 fibroblasts, a common model forcell-seeding in regenerative medicine. The same scheme of photochemicalattachment of propargyl-benzophenone followed by CuAAC reaction wasfollowed to introduce the RGD-azide onto the surface of the PCL fibers.The attachment of the azido-peptide was confirmed via X-rayphotoelectron spectroscopy (XPS), where the modified fiber showed asignificant nitrogen peak (N_(1s)) as would be expected for peptidemodified fibers, while no nitrogen peak was observed upon analysis ofthe PCL or PCL-benzophenone (FIG. 10B). XPS surface analysis indicatedan approximate surface coverage of 2% by mass, consistent with ourresults using the fluorescent azide.

PCL-RGD nanofibers were used as a cell seeding scaffold that would beable to promote adhesion, growth, and proliferation of NIH 3T3fibroblasts. Both PCL and PCL-RGD fibers were immobilized on a glassslide and NIH 3T3 cells were deposited onto the fiber. Following 72hours of incubation, cells were fixed, permeabilized, and stained usingactin green and DAPI. The slides were visualized via fluorescentconfocal microscopy (FIG. 10). After 72 hours, a much greater portion ofthe cells adhered to the fibers, as visualized by confocal microscopy.Additionally, the RGD-immobilized fibers provided enhanced cellspreading, as viusualized by the actin filaments within the cells. InFIGS. 10C and 9D, at 72 hours post-seeding, the cells have becomeelongated along the axis of the fiber and their proliferation wasenhanced by visual inspection of confocal micrographs. The unmodifiedfibers showed very little adhesion and proliferation after 72 hours. Toquantify proliferation of the cells on the fiber scaffold, a cellviability assay was used (i.e., MTT). After 72 hours of incubation, thefibers were removed from the slides and immersed in MTT containingmedia. The PCL-RGD fibers showed an approximate increase of 60%,relative to the PCL fibers alone after 72 hours of incubation. Theseresults indicate that the PCL-RGD fibers maintained the biologicalactivity of the peptide in sufficient concentrations to promoteadhesion, elongation, and proliferation.

This example shows the chemical modification of a continuously processednanofibrous biomaterial comprised solely from commodity polymers. Theprocessing technique is solvent free, scalable, uses FDA-friendlypolymers and allows for the simple tuning of cross-sectional dimensionsof the fiber. To convert this scaffold into a biological material, weutilized photochemistry to introduce an alkyne on to the surface of thefibers. This functional group allows for the modular synthesis of a hostof chemically derivatized fibers by employing the CuAAC reaction. Adensely covered surface was obtained using this technique, and moreimportantly the biological activity of the RGD peptide remained intactpromoting cellular adhesion and spreading.

Example 2

In this Example, we show the incorporation of a gradient of covalentlyimmobilized IKVAV peptides on coextruded PCL fibers to induce neuraldifferentiation and alignment. Coextruded fibers were chosen due totheir extensional strength, while maintaining lateral flexibility, in aneffort to mimic a spinal cord replacement. Additionally, extruded fiberscan remain aligned, providing for an additional topographical cue topromote directed elongation of neural cells. Photochemistry is used as aversatile tool to manipulate surface gradients, using a simple photomaskwith aligned PCL fibers. After attaching IKVAV gradients, we investigateneural cell growth in response to the gradient. In our system, bothgradient modification and fiber alignment dictate neural differentiationand cellular alignment.

Materials

N, N-dimethylformamide (DMF) (99%), dimethyl sulfoxide (DMSO), methanol(99.8%), Azide-fluor 488 (HPLC), 5-bromovaleric acid, sodium azide,trifluoroacetic acid (TFA) were purchased from Sigma Aldrich. Propargylbenzophenone was prepared as previously reported.Tris(3-hydroxypropyltriazolylmethyl)amine (THPTA) was a generous giftfrom the Finn lab. PC-12 adherent cells were purchased from ATCC. NerveGrowth Factor 7S was purchased from Life Technologies. 4′,6-diamidino-2-phenylindole (DAPI) was purchased from EMD Biosciences,Inc. Anti-Neuron-specific β-III Tubulin-NL557 was purchased from R&DSystems Inc.

Instrumentation

Multilayer co-extrusion was performed using two-component coextrusionsystem with 12 multipliers. ATR-FTIR Imaging was conducted on a DigilabFTS 7000 spectrometer, a UMA 600 microscope, and a 32×32 MCT IR Imagingfocal plane array (MCT-FPA) image detector with an average spatial areaof 176 μm×176 μm in the reflectance mode. Surface analysis of materialswas investigated on a PHI Versaprobe 5000 Scanning X-Ray PhotoelectronSpectrometer (XPS) with an Al Kα X-ray source (1486.6 eV photons).Scanning electron microscopy (SEM) was performed using a JEOL SEM underan emission voltage of 20 kV. A high-intensity UV lamp (Bluepoint 4Ecocure from Honle UV America Inc.) was used for surface modification ofthe PCL fibers with propargyl benzophenone (Pr-Bz). The molecular weightof the synthesized azido-peptide was measured on a Bruker AUTOFLEX IIIMALDI-TOF/TOF mass spectrometer using α-Cyano-4-hydroxycinnamic acid(CHCA) as a matrix. Fluorescent images were taken via laser scanningfluorescence confocal microscopy using a Leica TCS SPE ConfocalMicroscope. Water contact angle (WCA) measurements were tested on a CAM200 optical contact angle meter (KSV Instruments Ltd). Fluorescentgradient images were collected on a Maestro imaging system fromPerkinElmer.

Coextrusion of PCL Fibers

A multilayered film was extruded by a multiplication coextrusion processto fabricate polymeric fibers. PCL (CAPA 6800 pellets, MW=80 kg/mol) wascoextruded with poly(ethylene oxide) (PEO) to produce PCL fibers. Inorder to match the rheology of PCL and PEO melts during the extrusion,two grades of PEO (Dow POLYOX N80 (MW=200 kg/mol) and POLYOX N10 (MW=100kg/mol)) with a weight ratio of 30:70 were pre-blended using a HaakeRheodrive 5000 twin screw extruder. The viscosities of the obtained PEOblend and PCL melt match at the extrusion temperature, 200° C. Tenvertical multipliers and two horizontal multipliers were used throughoutthe extrusion line to generate a 256×4 matrix architecture that contains128×4 PCL domains embedded in PEO. The chill roll speed was 40 rpm andthe dimensions of the exit die are 0.5″×0.02″. PEO was removed bystirring in water at room temperature for 24 hours, to yield PCL fiberbundles. The PCL fibers were washed with methanol 3 times and vacuumdried overnight.

Photochemistry

10 mm×30 mm linear gradient images were printed on transparency films(3M PP2410/100) with a commercialized inkjet printer (Epson WF 3540)from transparent to black. To evaluate the gradient photomask, threenon-gradient photomasks were produced, corresponding to 3 points (black,50% black and transparent: 0, 1.5 and 3 cm on the gradient photomask) onthe gradient photomask to determine surface immobilization at each pointon the fiber (FIG. 11).

Each fiber bundle (3 cm, derived from a single tape) was soaked in aPr-Bz solution for 5 mins (10 mg/mL in methanol) and was air-dried atroom temperature. Samples were placed on a 25 cm×75 cm slide glass andcovered with the gradient photomask. The distance between the sample andUV source is 11.5 cm and a 320-390 nm filter was used. The UV intensityvaried linearly from 2.7 to 28.6 mW/cm² from black to transparent andthe center was 14.6 mW/cm². The unmasked UV source was 33.5 mW/cm².Samples were irradiated for 20 mins. After irradiation, samples werewashed in methanol overnight to remove excess Pr-Bz, and solvent wasevaporated under reduced pressure overnight. The process ofphotochemistry for non-gradients (PCL-ng-IKVAV) was repeated asdescribed above without the use of a photomask.

Click Chemistry

3 cm of PCL fiber bundles were immersed in a pre-mixed aqueous solutionof Azide Fluor 488 (AF₄₈₈) (0.8 mL, 3 mM) or azido-IKVAV (1.5 mL, 1.5mM), CuSO₄ (10 μL, 50 μM) and THPTA (50 μL, 50 μM). A fresh solution ofsodium ascorbate (100 μL, 100 mM) and 500 μL of DI water were added andincubated for 2 hours at room temperature. Gradient modified AF₄₈₈ orIKVAV (PCL-g-AF₄₈₈ or PCL-g-IKVAV) fibers were washed with dimethylsulfoxide (DMSO) or water, respectively, overnight to remove unreacteddye or peptide. The gradient conjugation of AF₄₈₈ with PCL-Alkyne fiberswas confirmed by Maestro fluorescence imaging (green excitation filterused with exposure time 100 ms). Quantification of surface AF488 dye wasdetermined by dissolution of the fibers in DCM and quantification byUV-Vis absorbance at 501 nm (FIG. 12). This value was compared to theBET surface area of the fibers to determine surface coverage density(FIG. 13). PCL-g-IKVAV fibers were scanned at 3 spots using ATR-FTIRcollecting from 600 to 4000 cm⁻¹ with 128 scans at the resolution 4cm⁻¹. Spatial distribution of absorbance at 1628 cm⁻¹ was taken usingFT-IR Imaging. Atomic chemical composition of PCL-g-IKVAV fibers wasanalyzed by XPS.

Wettability Test

Measurements were carried out at room temperature with high purity 18.2MΩ DI water. A single liquid droplet was suspended at the edge of asyringe needle (Matrix Technologies micro-Electrapette 25) and placed onthe measured surface. Water droplet images were taken on four differentspots across the PCL-g-IKVAV fibers for each sample. The contact angleswere measured as right and left angle of the water drop on the surfaceof the sample and reported as mean angles.

Cell Culture

PC-12 adherent cells (ATCC CRL-1721.1™) were purchased from ATCC andcells were cultured in F-12K Medium containing 15% heat inactivatedhorse serum, 2.5% fetal bovine serum (FBS), and 1% penicillin. Cellswere incubated in 75 cm² cell culture flasks at 37° C., in a 95% air and5% CO₂ environment. At 80-90% confluency, the cells were detached withPBS/EDTA for 10 minutes at 37° C. The detached cells were collected bycentrifugation at 800 g for 5 minutes. 14.5×10⁴ cells were placed oneach samples (PCL, PCL-ng-IKVAV and PCL-g-IKVAV) and cultured in thesame media with the addition of 50 ng/mL of NGF at 37° C., 5% CO₂ in ahumid environment for 5 days. The cell culture medium was refreshedevery 2 days.

Immunocytochemical Staining

After 5 days of incubation, all samples were washed with DPBS 3 timesand cells were fixed with 4% paraformaldehyde in DPBS for 15 minutes atroom temperature. After fixing cells, the samples were washed 3 timeswith DPBS. Non-specific binding sites were blocked with 10% normaldonkey serum, 0.1% Triton® X-100, and 1% BSA in DPBS for 30 minutes atroom temperature. Sequentially, cells were incubated withAnti-Neuron-specific β-III Tubulin NL557-conjugated antibody (1:10dilution) in blocking buffer for 3 hours at room temperature. After 3hours, cells were washed and nuclei were stained with DAPI (100 μL, 5μg/ml in PBS) to stain nuclei for 10 minutes and washed 3 times withDPBS.

Synthesis of IKVAV Peptide

The following is a representative coupling cycle for one amino acidmonomer—synthesis was carried out in a Torviq 25 mL filtered syringe,specifically designed for peptide synthesis. Peptide synthesis startedfrom 0.5 g of Rink Amide MBHA resin (0.52 mmol/gram) and this resin waspre-swelled in 15 mL of DMF for 10 minutes. The solvent is expelledthrough the syringe and a solution of 20% 4-methyl piperidine in DMF isadded to the resin (10 mL), this is carried out twice for 5 minutes and20 minutes, respectively. A Kaiser test was performed to confirm Fmocdeprotection. After a positive Kaiser test, the amino acid (0.78 mmol, 3equiv), HCTU (323 mg, 3 equiv) and DIPEA (272 μL, 6 equiv) weredissolved in a minimal amount of DMF. This solution was added to theresin at room temperature on a rotary shaker and allowed to react for 1hour. Following the coupling, the resin was again washed with DMF andDCM for 5 mins, 3 times each, and a qualitative Kaiser test wasperformed to ensure coupling. This synthetic process was repeated untilthe full peptide sequence was completed. After synthesis of the fullpeptide (IKVAV), the N-terminus of the peptide was conjugated with5-Bromovaleric acid—as described above. After washing 3 times with DMF(10 mL) and DCM (10 mL), sodium azide (85 mmol, 5 equiv) was added inDMF and allowed to react overnight. Subsequently, the resin was washedwith water to remove excess sodium azide and the resin was driedcompletely in a lyophilizer. To cleave the peptide, a solution ofTFA/H₂O (95/5, v/v) (10 mL) was incubated with the resin for 3 hours atroom temperature and the resin was removed by filtration. The cleavedpeptide solution was precipitated in cold ether, centrifuged (10,000 rpmfor 10 minutes at 4° C.), and decanted to yield a white pellet. Thefinal crude peptide was dried under vacuum to yield a white powder. Thecrude IKVAV peptides were purified by reverse phase-HPLC (LC-20AD pump,UV-VIS detector SPD-20A, Shimadzu HPLC System). Solution A is 0.05% TFAin water and solution B is 0.05% TFA in acetonitrile. Typical elutiongradients were performed using a C18 column (column size: 21.2×250 mm, 7μm particle size, Agilent ZORBAX 300SB-C18 PrepHT) using a lineargradient from 5 to 95% Solution B over 45 minutes. Collected fractionswere pooled and lyophilized to generate a white powder. Azido-IKVAVpeptide was confirmed via MALDI-TOF (positive ion mode) usingα-cyano-4-hydroxycinnamic acid (CHCA) as a matrix.

Surface Area Measurements

To measure the surface area of PCL fibers, BET measurements wereconducted. The samples were degassed with nitrogen gas to removeimpurities from the samples at 40° C. overnight. Krypton gas wasabsorbed on the surface of PCL fibers and 13 data points were collectedwith the relative pressure ranging from 0.06 to 0.30. The BET plot wasgenerated with a linear region that has a slope of 2.773±0.048 g/cm³ andan intercept of 0.228±0.010 g/cm³ (R²=0.998).

Results

Coextruded Aligned PCL Fiber

In this example, multilayered melt coextrusion of polymeric fibers waschosen for several reasons. The layered coextrusion system uses twopolymer components, in this case PCL and PEO, which are both polymersthat have been used in several FDA-approved applications. Additionally,both polymers are inexpensive, costing less than $15/kg. Finally, themethod is continuous and solvent free, requiring only a water wash togenerate PCL fibers, hence eliminating biologically detrimental effectsof solvent based processing. During the extrusion process, PCL and poly(ethylene oxide) (PEO) are melt-pumped and layered vertically in theextrusion line. This vertical bilayer was cut in a horizontal plane inthe multiplication die and recombined side-by-side to double the numberof vertical layers. This process was repeated ten times to increase thenumber of layers and conversely, decrease the layer thickness. Followingvertical multiplication, PEO is layered on the top and bottom of thevertical multilayers. Finally, the multilayered melt undergoeshorizontal multiplication, where it is cut in the vertical direction,and the two flow fields are stacked on top of each other to yield PCLfibers embedded in a PEO tape. This procedure was repeated two times,producing a composite extrudate tape containing 512 PCL domains embeddedin a PEO tape. The extruded tape is washed in a water bath for ˜24 hoursto remove PEO, yielding PCL fibers. Distribution of the size of PCLfibers was examined using scanning electron microscopy (SEM) and wasanalyzed. The fiber dimensions shows a narrow distribution of size withan average cross-section of 1.49±0.48 μm.

Gradient Photochemistry

In this example, we sought to create a simple method to generatesurface-immobilized chemical gradients on aligned extruded fibers. Owingto the ease with which photochemical modification occurs, we aimed toutilize a simple photomasking strategy to modulate the concentration offunctionalized benzophenone deposited on the fibers. A 3 cm linearphotomask was printed on transparency slides ranging from completelytransparent to entirely black (FIG. 12A), allowing us to modulate theUV-intensity for photochemistry by one full order of magnitude. PCLfibers were dip coated in a concentrated solution of propargylbenzophenone (Pr-Bz), allowed to dry, and subjected to UV-irradiationfor 20 minutes (FIG. 11).

Upon UV irradiation, Pr-Bz is excited to form a radical species, whichcan then undergo a radical insertion into the PCL backbone. Thisresulted in a new covalent bond between Pr-Bz and PCL, leaving surfaceexposed propargyl groups that could undergo the CuAAC reaction. Afteralkyne gradient formation, the CuAAC reaction was used to attach AzideFluor 488 (AF₄₈₈) to the fibers, a green fluorescent dye that allows forsimple visualization of the gradient modified surface. The surfacegradient was visualized using fluorescence imaging in the green channel,clearly indicating gradient formation (FIG. 12B). The gradient intensitywas analyzed by mean fluorescence intensity over the entire fiber bundleto correlate the fluorescent gradient to the linear photomask, showing astrong linear correlation to the mask (FIG. 12C).

Quantification of AF₄₈₈ Gradient

In order to quantify surface coverage following AF₄₈₈ immobilization,dye coverage was quantified using UV-Vis spectroscopy. Dyequantification proves to be much simpler than peptide quantification,and in the past, has correlated well to peptide conjugation. Thecorrelation is likely due to the extreme efficiency of theligand-accelerated CuAAC reaction. Rather than quantifying the gradientfibers directly, we chose to take individual points within the gradientand understand how UV fluence impacted surface coverage. Threeindividual photomasks were used to attach Pr-Bz, followed by CuAAC withAF₄₈₈, where photomasks corresponded to (a), (b), and (c) in FIG. 12A(i.e., completely transparent, 50% black, and 100% black) to evaluatesurface coverage (FIG. 16). After the CuAAC reaction, fibers weredissolved and dye loading was quantified and compared to BET surfacearea measurements (18.8 cm²/mg). The quantification of AF₄₈₈ wasdetermined using UV-Vis spectroscopy at 501 nm against a standard curvein dichloromethane following dissolution of PCL-AF488 fibers. Fiberswith a transparent photomask, corresponding to spot (c) (FIGS. 12A and16), allowed UV transmittance of 28.6 mW/cm² and led to 0.43 nM/cm² ofsurface coverage (FIG. 17) on the fiber bundle. An intermediateintensity, corresponding to 50% black in the photomask indicated by spot(b) (FIG. 12A) provided 14.6 mW/cm² of UV intensity. This photomaskyielded 0.24 nM/cm² of AF₄₈₈ attachment onto the surface of the fibers(FIG. 17). Finally, the UV intensity of the 100% black photomask,corresponding to spot (a) (FIG. 12A) of the gradient photomask yielded0.09 nM/cm² of AF₄₈₈ decorated onto the fibers. Gratifyingly, the simpleinkjet photomasking technique yielded approximate linear results thatallowed us to tune the gradients by nearly an order of magnitude from0.43 to 0.09 nM/cm² onto the surface.

Gradient Surface Modification of PCL with IKVAV

It is well known that cells can respond to haptotactic gradients, orsurface immobilized gradients of biologically active molecules. Suchgradients can lead to enhanced cell adhesion and migration relative tonon-gradient surfaces. In particular, neuronal cells are especiallysensitive to these chemical perturbations and can undergo axonalelongation in the presence of haptotactic gradients of laminin derivedpeptides, like IKVAV of YIGSR. We sought to employ this phenomenon withour extruded fibers, employing the same linear gradient as describedabove. An azide-modified IKVAV was synthesized and clicked onto Pr-Bzmodified PCL fibers, using the optimized gradient conditions forphotochemistry and ligand-accelerated CuAAC (PCL-g-IKVAV). As IKVAV doesnot provide the ease of quantification of the dye-immobilized molecules,several characterization techniques were carried out to investigatepeptide gradient formation. First, PCL-g-IKVAV was analyzed by ATR-FTIRand FTIR imaging. The C═O stretching mode in the amide I region iscoupled to the bending of the N—H bond and the stretching of the C—Nbond represented between 1620 and 1640 cm⁻¹. The ATR-FTIR spectra at1628 cm⁻¹ confirmed an increasing amide I band (C═O) across the breadthof the fiber length of PCL-g-IKVAV with detection at the 3 differentregions of the sample (FIG. 13B). With increasing transparencies of thephotomask, more intense amide I peaks are seen, indicating an increasinggradient of amide bonds, as would be expected for gradient IKVAVformation. FTIR imaging was also employed to visualize the spatialdistribution of the C═O amide I band and is shown as a chemical heat map(FIG. 13C) from spots (a) to (c) (scale bar on the right indicatesintensity of absorbance, blue to red after normalization). The FTIRimaging result correlates well with the full ATR-FTIR spectra as anincrease in the intensity of amide C═O bonds is seen, indicating anincrease in concentration of IKVAV.

Water contact angle provides an indication of the hydrophilicity andhydrophobicity on the surface of polymeric substrates. Cell adhesionalso depends on the wettability of a scaffold surface, especially sincesynthetic biocompatible polymers, such as PCL, are hydrophobic whichusually limits cell interactions with the scaffold. Incorporatingpeptides such as IKVAV or YIGSR will likely improve the hydrophilicityof the surface, in addition to interacting with cell surface receptors,further enhancing the cell adhesion properties of the polymericscaffold. Water contact angles were measured on PCL-g-IKVAV fibers fromspot 1 to spot 4 indicating a decreasing water contact angle, and hencean increasing amount of surface-immobilized IKVAV (FIG. 14A). Watercontact angle values decreased from 107.8±7.5 to 65.5±3.7° (FIG. 14B),indicating that PCL-g-IKVAV fibers become more hydrophilic as surfacedensity is increased. Furthermore, this follows a linear trend as wouldbe expected from our linear photomask. Therefore, it is likely that bothgradient hydrophilicity and receptor specific interactions may play arole in improving cell surface adhesion.

Finally, PCL-g-IKVAV fibers were characterized by X-ray photoelectronspectroscopy (XPS) to determine nitrogen content, a unique atom afterthe fibers are peptide-modified. XPS wide scan of PCL-g-IKVAV fibers,(FIG. 14C) was performed at 4 distinct spots at increasing gradientdensities, and the intensity of the N1s was quantified. An increasingintensity of N1s (relative %) at −400 eV was seen, showing successfulimmobilization of gradiated amounts of IKVAV onto the PCL fibers. Thepercentages of nitrogen were 3.5, 7.5, 9.8 and 12.0% corresponding tospot 1 to 4 (FIG. 15C) as a result of gradient IKVAV immobilization (seeTable 1). This result also shows the linear behavior of the nitrogen tocarbon ratio (N/C), confirming linear gradient formation.

TABLE 1 Surface characterization - Relative elemental concentrations ofdifferent spots on the PCL-g-IKVAV fibers by XPS and values for watercontact angles (right column) Atomic component (%) Sample C (1s) N(1s) O(1s) WCA angle (°) Spot 1 77.9 3.5 18.6 107.8 ± 7.5  Spot 2 71.7 7.520.7 89.8 ± 3.9 Spot 3 70.4 9.8 19.8 74.6 ± 8.1 Spot 4 70.1 12.0 18.065.5 ± 3.7In Vitro Characterization of Aligned PCL-g-IKVAV Fibers with PC-12 Cells

To determine the effects of the gradient on neural cells, growth ofPC-12 cells was evaluated on three different substrates, unmodified PCLfibers, PCL non-gradient IKVAV (PCL-ng-IKVAV) and PCL-g-IKVAV. PC-12cells are derived from a pheochromocytoma of the rat adrenal medulla,which can differentiate into neurons and are commonly used as a modelcell line in regenerative neural medicine. PC-12 cells were seeded ontothe fiber and cultured for 5 days in the presence of nerve growth factor(NGF) to allow for neural differentiation and neurite extension.Following incubation, cells were fixed and stained using a NL557conjugated anti β-III-tubulin antibody that is specific for neuraldifferentiation (red) and DAPI for the nucleus (blue) (FIG. 15).Confocal microscopy images reveal different neurite extension andcellular density of PC-12 cells on each substrate, as well as in threedifferent regions of each substrate. The density of cells on theunmodified PCL fibers, was significantly lower than that observed on thePCL-ng-IKVAV and PCL-g-IKVAV (FIG. 5). Also the 3 different areas on theunmodified PCL showed little adhesion, differentiation, and directedcell alignment (FIGS. 15A-C). This would be as expected, as PCL providesno biologically active cues and the hydrophobic nature of the surfaceprovides little adhesive capabilities for cells.

The biologically active peptide, IKVAV, is known to bind to β-amyloidprecursor protein and has been shown to promote adhesion and neuriteoutgrowth of PC-12 cells. It was postulated that PCL-ng-IKVAV would leadto improved adhesion but would have less propensity for cell alignmentand extension than PCL-g-IKVAV fibers. Confocal images indicate a higherdensity of cells on non-gradient substrates as higher IKVAV densitiesare seen throughout the non-gradient fibers (FIGS. 15D-F). Althoughhigher IKVAV densities enhanced cell adhesion on PCL-ng-IKVAV, there isminimal neuronal alignment or differentiation of PC-12 cells with afixed concentration of IKVAV across the sample. Spreading of neuriteswas in arbitrary directions in the three different spots as observed viaanti β-III-tubulin, and neural differentiation was less significant whencompared to gradient modified fibers. When non-gradient substrates arecompared to PCL-g-IKVAV, several differences arise. FIG. 15G,corresponds to the lowest concentration of IKVAV on the gradient andshows lower numbers of PC-12 cells, as compared to other regions on thesame substrate. The highest cell density was seen on the highestconcentrations of IKVAV on the gradient fibers (FIG. 15I). In addition,PC-12 cells had noticeably elongated nuclei and neurites along the axisof the PCL-g-IKVAV fibers. The primary direction was along the gradientof the IKVAV motif, which showed the highest cell extension,differentiation and alignment of neurites, as compared to all othersamples (FIG. 15G-I). Regardless of peptide concentration on thegradient surface, neurite extension was seen both in the direction ofthe fibers and along the axis of the gradient. This result implies thatthe gradient surface of PCL-g-IKVAV fibers is critical for strongbiological activity and that the higher concentrations of IKVAV impactedcell adhesion. The gradient fibers effect differentiation of PC-12 cellsand their alignment more so than solely the topographical cues of thealigned fiber direction, although both components are interrelated.

Photochemical gradient modification of a scalable class of meltcoextruded fibers allowed for immobilization of azide-modified peptidegradients on the surface of aligned PCL fibers. Melt coextrusion allowsfor a significantly higher throughput of fiber production when comparedto other common fiber processing techniques. This photochemicalmodification is modular and allows for patterning of surface groupsabout the fiber. Gradient immobilization of the laminin derived peptide,IKVAV, onto PCL fibers was easily accomplished.

Example 3

We developed a wound healing patch where a genetically-engineered EGFwas covalently immobilized onto a polymeric fiber mat, thus eliminatingthe chemical driving force for release to the environment. AMMP-cleavage site was introduced between the active protein and thecovalent attachment point on the mat, thus EGF would be liberated byMMP-9, a commonly upregulated MMP during the early wound healing stages(FIG. 25). The protein-conjugated fiber mat is envisioned as a topicaldepot of EGF, which can be tuned to provide the optimal amounts of EGFfor treatment of acute and chronic wounds. An enzymatically responsiverecombinant EGF was synthesized with several features included at theN-terminus of the protein (EGF-MMP, FIG. 18A). The N-terminus wasmodified since the C-terminal sequence is responsible for receptorbinding. The recombinant protein contained a N-terminal sequence (AKT)that is highly reactive for bioconjugation using pyridoxal 5′-phosphate(PLP) mediated bioconjugation. This method is a site-selectivemodification that utilizes N-terminal transamination of proteins tointroduce a ketone or aldehyde, which could further undergo oximeligation with a polymer substrate (see below). Next a polyhistidine tagwas included for affinity purification. Finally, a MMP cleavage site fortriggered release of the protein from the scaffold was includeddownstream of the remaining N-terminal extension. The chosen octapeptidesequence (VPLSLYSG) (SEQ ID NO: 1) is cleavable by MMPs, including MMP-9(kcat/KM=49,000±3,000), and is selectively cleaved by MMPs that areupregulated in early stages of the wound healing cascade. Only a4-residue extension remains at the N-terminus following release, basedon this domain orientation.

The gene was synthesized and cloned into the pET28a(+) expression vector(pET28EGF-MMP), followed by standard IPTGinduced protein expression andpurified via affinity chromatography. An EGF mutant without theMMP-cleavage site was synthesized in the same manner as a non-responsivecontrol (FIG. 18B). The two proteins were analyzed via SDS-PAGE todetermine purity, resulting in a single band corresponding to amolecular weight of −10 kDa (FIG. 18C). MALDI-TOF mass spectrometryconfirmed the molecular weight of the recombinant proteins, which areconsistent with the theoretical molecular weights of control EGF andEGF-MMP, 7,344 kDa and 8,162 kDa (expected molecular weight 7,344 kDaand 8,162 kDa), respectively (FIG. 18D).

The in vitro biological activity of the purified EGF-MMP protein wasconfirmed using both human keratinocytes (HaCaT) and epidermoidcarcinoma (A431) cell lines. EGF is a potent mitogen in HaCaT cells,even at low concentrations, and improves further at high concentrations.EGF has the opposite biological effect on A431 cells, promotingapoptosis at high concentrations.28 Relative viability was evaluatedutilizing the MTT proliferation assay on the two cell lines. HaCaT cellsdemonstrated a ˜6-fold increase in cell viability from 100 to 587.7% atthe highest concentration of EGF-MMP (FIG. 19A). In contrast, the cellviability of A431 cells decreased from 100 to 38.2% as the concentrationof EGFMMP increased. These results were consistent with literaturevalues and demonstrated that the purified recombinant EGF-MMP has theexpected biological activity and the mutations introduced at theN-terminus do not affect the activity.

In silico modeling predicted that the MMP reactive octapeptide would becleaved by MMP-9 orders of magnitude faster than other amino acidsequences in EGF-MMP. Cleavage of EGFMMP was performed via incubationwith MMP-9 for 2 and 4 hours (FIG. 19B). The molecular weight of theprotein was determined by MALDI-TOF following cleavage. After 2 hours,the molecular weight decreased from 8.16 kDa to 6.64 kDa, in preciseagreement with the theoretical molecular weight of cleaved EGFMMP (6.64kDa). After 4 hours, the mass spectrum remained as a single peak at 6.64kDa, indicating no proteolytic activity at other sites on the protein.This result indicated over 99% conversion to the cleaved product byintegration of MALDI-TOF data, with no side products.

The PLP-mediated transamination reaction was employed to introduce aketone moiety at the N-terminus of EGF for conjugation to the fiber matand conversion was verified using MALDI-TOF. Two significant peaks werefound (FIG. 22), one corresponding to the transamination product andanother that correlates to an expected PLP adduct. The undesired productstill contained a ketone group and was able to undergo oxime formation.Bioorthogonal ligation has been widely explored in chemical biology forthe reaction of ketones with hydrazides and alkoxyamines to formhydrazones and oximes at acidic pH. Oxime chemistry prefers slightlyacidic conditions, however proteins may precipitate under theseconditions due to the relatively higher isoelectric points of proteins(pIEGF-MMP=6.2) than those that are preferable for oxime ligation.Aniline catalysts effectively accelerate oxime formation by forming anin situ highly reactive protonated aniline Schiff base at neutral pH.Therefore, aniline was utilized as a nucleophilic catalyst for thebioconjugation reaction. EGF-MMP was modified with the ketone moiety atthe Nterminus in the presence of PLP, as previously described. ExcessPLP was removed via centrifugal spin filtration and the ketonefunctionalized protein was conjugated to aminooxy functionalized TAMRAdye with a minimal amount of aniline catalyst (FIG. 23A). After removalof free dye and catalyst via centrifugal spin filtration, fast proteinliquid chromatography (FPLC) was performed to monitor the elution volumeof the PLP reacted protein and dye labeled EGF-MMP, both the modifiedand unmodified proteins showed identical elution volumes (FIG. 23B).Furthermore, after oxime ligation a peak at 555 nm co-eluted with theprotein peaks, demonstrating successful bioconjugation. The polymericmaterial chosen was a microfibrous coextruded poly(ε-caprolactone) (PCL)non-woven mat. Multilayered coextrusion of nano- and microfibers is anew manufacturing technique that is high throughput, allows for controlover fiber dimensions and porosity, and is amenable to site-specificchemical modification strategies. Non-woven poly(ε-caprolactone) (PCL)fiber mats were fabricated (FIG. 20A) with a high porosity and nearhomogeneous pore size distribution (25.6 μm, FIG. 20B). This pore sizeis in the range of commercially available wound healing patches. In thiswork, PCL was chosen due to its processing flexibility andbiocompatibility, specifically as a wound dressing.

Aminooxy benzophenone was synthesized as a linker for oxime chemistry,and deposited onto the PCL fiber mat through UVinitiated photochemistry.Ketone-modified EGF-MMP was then covalently conjugated onto the aminooxyfiber mat via oxime ligation. Since protein adsorption without covalentattachment can provide false readings, non-specific protein absorptiononto the hydrophobic fiber mat was performed on unmodified mats. Theprotein modified PCL fiber mat was scanned using ATRFTIR to confirm thepresence of EGF-MMP (FIG. 20C and FIG. 24). The un-irradiated fiber matshowed no amide signal from the protein, while the UV irradiated sideclearly showed EGF-MMP, as indicated by C═O stretching vibrations at1615 cm−1 from amide I, N—H bending vibration/C—N stretching vibrationat 1505 cm−1 from amide II, and N—H stretching vibration near 3300 cm−1from amide A. This result clearly indicated that protein conjugationonly proceeds onto the aminooxy modified PCL fiber mat. We next exploredwhether incubation with MMP-9 releases EGF conjugated to the fiber mat.For comparison, a conjugated noncleavable control EGF was also studied.The release kinetics of EGF-MMP and control EGF from the fiber mats weredetermined by incubation with activated MMP-9 and quantification viaELISA. The concentration of released protein was normalized by theweight of each fiber mat. The ELISA results indicated that EGF-MMP wasrapidly released, resulting in ˜11 ng EGF/mg PCL released over the first10 minutes after introduction of the enzyme (FIG. 20D). However, thecontrol EGF conjugated fiber mat showed no significant protein releaseafter introduction of the MMP enzyme. After 4 hours of incubation, thecontrol EGF protein showed a negligible amount of released protein whilethe EGF-MMP fiber mat released ˜20 ng of EGF per milligram of PCL fibermat. Incubation was carried out for 24 hours with MMP-9, with no furtherincrease in released protein. Thus, the conjugation scheme yielded 20 ngEGF/mg PCL that was accessible. When normalized per surface area, basedon Brunner-Emmett-Teller (BET) surface measurement (43.2 cm2/mg of PCLfiber mat), ˜0.46 ng EGF/cm2 of PCL fiber mat was available for release.The results indicated that EGF-MMP release was specifically triggered byMMP-9, while the control EGF remained conjugated to the polymerscaffold.

Wound healing was simulated using a monolayer scratch test with HaCaTcells, a common model for migration and proliferation in the woundhealing process (FIG. 21). A control sample with no fiber mat and threedifferent fiber mats were evaluated; unmodified PCL, controlEGF-modified PCL, and EGF-MMP-modified PCL. Cells were grown toconfluency and a scratch was introduced using a micropipette tip. Thecells were then incubated with a fiber mat and MMP-9. After 24 hours ofincubation, the scratches were measured by optical microscopy. Noscratch closure was observed for controls with neat PCL or no fiber mat(FIG. 21). We observed a decreased gap for cells incubated with controlEGF fiber mats. The improvement is likely due to a small amount of EGFreleased, either through non-specific adsorption to the surface orlow-level cleavage of unintended sites. Previous studies have shown EGFconcentrations as low as 1 ng/mL can have a pronounced effect on gapclosure. In contrast, the EGF-MMP mat demonstrated complete gap closureand significantly enhanced proliferation. Thus, the EGF-MMP fiber matresponded to MMP-9 to stimulate complete wound closure in the simulatedhealing.

This study describes a new paradigm in triggered release of growthfactors based on genetic engineering, where therapeutic protein releaseis triggered by the biology of wound healing. Recombinant EGF wasengineered to release only in the presence of wound healing cues. Theresults presented here clearly indicate that EGF could be engineered forselective release by MMP-9.

Moreover, released EGF from the EGF-MMP conjugated fiber promotedproliferation and migration of cells. The amount of EGF delivered inresponse to MMP-9 can be tuned in the future via the amount of aminooxygroups coupled onto the fiber mat and release kinetics can be controlledby changing the MMP responsive sequence.

Experimental Section

Materials

Aminooxy-5(6)-TAMRA was purchased from Biotium, Inc. BL21(DE3) E. coliwere purchased from Genlantis. Aniline,1,8-Diazabicyclo[5.4.0]undec-7-ene (DBU), N-hydroxyphthalimide, andα-cyano-4-hydroxycinnamic acid were purchased from Sigma-Aldrich. 1-StepTMB substrate, agarose, calcium chloride, isopropylβ-D-1-thiogalactopyranoside, kanamycin sulfate, potassium phosphatemonobasic anhydrous, potassium phosphate dibasic anhydrous, pyridoxal5′-phosphate, Miller LB broth, methanol, SeeBlue Plus2 protein ladder,sodium hydroxide, sulfuric acid, Tween-20, Tris-HCl, potassiumcarbonate, N,N-dimethylformamide, potassium iodine, sodium bicarbonate,chloroform-d6, and urea were obtained from Fisher Scientific.Recombinant, human pro-MMP-9 expressed in CHO cells and MMP-2/MMP-9Substrate I, Fluorogenic were purchased from EMD Millipore. Dimethylsulfoxide (DMSO) was purchased from Amresco. Amino-phenyl mercuricacetate, imidazole, 4-hydroxyl benzophenone, 2-bromoethanol,dichloromethane, trimethylamine, and Triton X-114 were purchased fromAcros Organics. Chemically competent NEB 5 α E. coli, NcoI restrictionenzyme, XhoI restriction enzyme, T4 polynucleotide kinase, and T4 ligasewere purchased from New England Biolabs. Recombinant human epidermalgrowth factor was purchased from BD Bioscience. Mouse anti-humanepidermal growth factor IgG and horse radish peroxidase conjugated goatanti-mouse IgG secondary antibody was purchased from Life Technologies.HaCaT and A431 cells were generous gifts from the dermatology departmentat Case Western Reserve University.

Instrumentation

Proton nuclear magnetic resonance (¹H NMR) were recorded on a VarianInova 600 MHz NMR spectrometer in deuterated solvents. Chemical shiftsare reported in parts per million (ppm, δ) relative to residual solvent(CDCl3, δ 7.26). Fast protein liquid chromatography (FPLC) was performedusing a GE Healthcare AKTAFPLC 900 chromatography system equipped with aSuperdex 75 10/300 GL size exclusion column. A Thermo Finnigan LCQAdvantage LC/MS (ESI) was used to confirm the molecular weight ofsynthesized aminooxy benzophenone. SDS polyacrylamide gelelectrophoresis (PAGE) was performed on Novex NuPAGE 4-12% bis-trisprotein gels (1.0 mm×12 well) (35 minutes, 200 V, 1× NuPAGE MES SDSrunning buffer). Gels were stained with Coomassie SimplyBlue SafeStain(Life Technologies). Multilayer coextrusion was performed using theCLiPS two-component coextrusion system with 23 multipliers. ATR-FTIRimaging was conducted on a Digilab FTS 7000 spectrometer, a UMA 600microscope. A high-intensity UV lamp (Bluepoint 4 Ecocure from Honle UVAmerica Inc.) was used for surface modification of the PCL fibers withaminooxy benzophenone. The molecular weights of the synthesized proteinswere measured on a Bruker Autoflex III MALDI-TOF/TOF mass spectrometerusing α-cyano-4-hydroxycinnamic acid (CHCA) as a matrix. UV-vis spectrawere collected using a Shimadzu BioSpecnano UV-vis spectrophotometer.

Synthesis of Aminooxy-Benzophenone

Aminooxy-benzophenone was synthesized. 2-(4-Benzoylphenoxy)ethanol (1).4-Hydroxylbenzophenone (2 g, 10 mmol) and anhydrous potassium carbonate(2.76 g, 20 mmol) were stirred in a round bottom flask in 30 mL ofN,N-dimethylformamide (DMF). 2-Bromoethanol (1.07 mL, 15 mmol) andpotassium iodide (KI) (0.8 g, 4.8 mmol) were added, and the reactionmixture was heated for 24 hours at 65° C. After the reaction, themixture was cooled to room temperature and then filtered. The yellowfiltrate was slowly precipitated in 700 mL of deionized water in an icebath. A white powder was collected by centrifuging at 10,000 rpm for 10min (80% yield) 1H NMR (600 MHz, chloroform-d) δ 7.82 (d, 2H), 7.76 (d,2H), 7.57 (t, 1H), 7.47 (t, 2H), 6.99 (d, 2H) 4.17 (t, 2H), 4.01 (t,2H). ESI-MS (m/z, rel %) 243.0 ([M+H]+, 10%), 264.9 ([M+Na]+, 90%)calculated for C₁₅H₁₄O₃.

Methyl sulfonyl benzophenone (2). 2-(4-Benzoylphenoxy)ethanol (2 g, 8mmol) in DCM was cooled to 0° C. in an icebath. Triethylamine (TEA) (2g, 2 mmol) was mixed and then methyl sulfonyl chloride (2.27 g, 1.2mmol) was added by syringe into the same flask. The reaction wasperformed at room temperature and stirred overnight in a nitrogenatmosphere. Crude material was sequentially washed with saturated NaHCO₃and brine solution. The organic layer was dried in vacuo. ¹H NMR (600MHz, chloroform-d) δ 7.84 (d, 2H), 7.75 (d, 2H), 7.57 (t, 1H), 7.48 (t,2H), 6.98 (d, 2H), 4.61 (t, 2H), 4.33 (t, 2H), 3.1 (s, 3H) ppm. ESI-MS(m/z, rel %) 321.0 ([M+H]+, 100%), calculated for C₁₆H₁₆O₅S.

Aminooxy benzophenone (4). Methyl sulfonyl benzophenone (2 g, 6 mmol)was added to a round bottomed flask with1,8-diazabicyclo[5.4.0]undec-7-ene (DBU) (0.93 mL, 7.5 mmol) andN-hydroxyphthalimide (1.3 g, 7.5 mmol) in DMF (50 mL). The reactionproceeded overnight at room temperature with stirring. The followingday, DMF was removed by rotary evaporation. The crude reaction wasre-suspended in dichloromethane (DCM) and DBU was filtered via flashsilica column chromatography. The filtrate in DCM was washed with brineand sodium bicarbonate, dried over anhydrous sodium sulfate, andfiltered. The dried phthalimide benzophenone (3) (1 g, 2.6 mmol) wasre-dissolved in acetonitrile (50 mL). Hydrazine monohydrate (0.42 g, 13mmol) was added and the mixture was stirred for two hours at roomtemperature. After concentrating the reaction by rotary evaporation, 20mL of DCM was added and the mixture was filtered over a plug of celiteunder vacuum. The product was purified via silica flash chromatography(n-hexane:ethyl acetate, 1:2, v/v). ¹H NMR (600 MHz, chloroform-d) δ7.88 (d, 2H), 7.79 (d, 2H), 7.61 (t, 1H), 7.52 (t, 2H), 7.02 (d, 2H),5.60 (s, 2H) 4.28 (t, 2H), 4.02 (t, 2H) ppm. ESI-MS (m/z, rel %) 280.0([M+Na]+, 100%), calculated for C₁₅H₁₅NO₃.

Protein Sequences and Plasmid Generation

Epidermal growth factor used in this study was produced recombinantly inE. coli to incorporate a PLP reactive site and MMP cleavage site intothe sequence. The protein sequence for the mutant, referred to asEGF-MMP, is as follows:

(SEQ ID NO: 2) AKTHHHHHHVPLSLYSGNSDSECPLSHDGYCLHDGVCMYIEALDKYACNCVVGYIGERCQYRDLKWWELR

The DNA sequence was generated from the protein sequence usingGeneDesign 2.0 software and optimized for E. coli codon usage. The DNAsequence is given below with the added NcoI and XhoI cut sitesunderlined at the 5′ and 3′ ends respectively.

(SEQ ID NO: 3) 5′ ATGCTACCATGGCTAAAACCCACCACCACCACCACCACGTTCCGCTGTCTCTGTACTCTGGTAACTCTGACTCTGAATGCCCGCTGTCTCACGACGGTTACTGCCTGCACGACGGTGTTTGCATGTACATCGAAGCTCTGGACAAATACGCTTGCAACTGCGTTGTTGGTTACATCGGTGAACGTTGCCAGTACCGTGACCTGAAATGGTGGGAACTGCGTTAACTCGAGTGACTC 3′

The gene was produced via primer overlap PCR of primers purchased fromIntegrated DNA Technologies designed by GeneDesign 2.0 software. The PCRwas run for 55 cycles and the product was purified using an agarose gel,the band excised, and the excised band was spin column purified. Thepurified product was digested with NcoI and XhoI and spin columnpurified. The product was then ligated via T7 DNA ligase into apET28a(+) vector that had been digested with NcoI and XhoI. The ligationmixture was transformed into chemically competent NEB5α E. coli andplasmids were extracted from isolated colonies. The insertion of thesequence into the plasmid was verified via sequencing with T7 promoterprimers and the pET28(EGF-MMP) plasmid was isolated. A mutant of theEGF-MMP protein was made to remove the MMP cutsite for controlexperiments (referred to as EGF), with the protein sequence given below:

(SEQ ID NO: 4) AKTHHHHHHNSDSECPLSHDGYCLHDGVCMYIEALDKYACNCVVGYIGERCQYRDLKWWELR

The PLP reactive site is highlighted in red and the His tag forpurification is highlighted in green. Primers were designed forwhole-plasmid PCR of the pET28(EGF-MMP) plasmid with the removal of theDNA sequence encoding the MMP cut-site amino acid sequence. PCR was runwith the primers and pET28(EGF-MMP) plasmid and the resulting PCRmixture was treated with T7 polynucleotide kinase and then ligated withT7 DNA polymerase. The ligation mixture was transformed into chemicallycompetent NEB5α E. coli cells and isolated colonies had plasmidssequenced as previously described. The pET28(EGF) plasmid was isolatedfrom successfully transformed colonies.

Recombinant EGF Protein Expression and Purification

Chemically competent BL21(DE3) E. coli cells were transformed withpET28(EGF-MMP) or pET28(EGF) and plated onto LB agar media containing 50μg/mL kanamycin. After overnight incubation at 37° C., 100 mL of LBmedia containing 50 μg/mL kanamycin was inoculated with a well-isolatedcolony and incubated overnight at 37° C. with agitation at 250 rpm. Theovernight culture was then diluted into 1000 mL of LB media containing50 μg/mL kanamycin and incubated at 37° C. with agitation at 250 rpm.Cell culture growth was monitored by optical density at 600 nm (OD600)utilizing UV-vis spectroscopy. When the OD600 of the cultures reachedapproximately 0.8 (mid-log phase), protein expression was induced byaddition of 10 mL of 100 mM IPTG resulting in a final concentration of 1mM. Shaking was continued at 37° C. for an additional 6 hours, at whichpoint cells were collected by centrifugation in an Eppendorf A-4-81rotor at 4000 rpm (4° C.) for 30 min. The supernatant was decanted andthe cell pellet was frozen at −80° C. overnight. Cells were resuspendedin 50 mL of 50 mM phosphate buffer with 500 mM NaCl (pH 7.4) and thisbuffer was used for subsequent steps in the purification. Samples werelysed with a microtip sonicator (10 min total sonication time, cycles of30 s on/30 s off, power output of 6) in an ice bath. The resulting celldebris was pelleted in an Eppendorf A-4-81 rotor at 4000 rpm (4° C.) for30 min and the supernatant was decanted. The expressed protein waspresent in the pellet as an inclusion body. The inclusion bodies weredenatured by incubating the re-suspended pellets in 50 mL of 4M ureaphosphate buffer for 1 hour at 4° C. Solids were pulled out of thesolution and the inclusion body solution was incubated with 8 mL ofHisPur cobalt resin that had been washed and equilibrated to buffer. Theinclusion body solution was allowed to incubate for 1 hour and thenloaded into a fritted syringe. The resin bed was allowed to settle undergravity flow and washed with 4 M urea buffer containing 5 mM imidazolefor 16 column volumes. The resin bed was then eluted with an imidazolegradient all made in 4 M urea buffer of 15, 25, 35, 45, 55, 65, 75, 85,95, and 105 mM imidazole. 6 mL was used for each concentration andfractions were collected under gravity flow. The column was fully elutedusing 6 mL of 150 mM imidazole and the fractions were analyzed usingSDS-PAGE. Fractions containing pure recombinant protein were pooledtogether and refolded through sequential dialysis against buffercontaining 3, 2, and 1 M urea using 3.5K MWCO dialysis tubing for aminimum of 4 hours each. The refolding was then completed via dialysisagainst phosphate buffer 3 times for a minimum of 4 hours each. Therefolded protein solution was concentrated using 3.5K MWCO ofcentrifugal spin filtration (8,000 rpm) and quantified via 280 nmabsorbance (ε280=20,315 and 18,825 M−1 cm−1 for EGF-MMP and EGFrespectively). The purified proteins were analyzed via SDS-PAGE, FPLC,MALDI-TOF MS.

EGF-MMP Cleavage Test by MMP-9

100 mg/mL of EGF-MMP was prepared (n=3) in 2 mL of PBS buffer. In orderto activate MMP-9 enzyme, amino-phenyl mercuric acetate (APMA) (3.5 mg,10 mmol) was dissolved in 1 ml of DMSO and the solution was diluted withreaction buffer (50 mM Tris-HCL pH 7.4, 1 mM CaCl2, 0.05% Triton X-100)to a final concentration of 2 mM APMA. 100 μL of the APMA solution wasadded to 900 μL of phosphate buffer (50 mM phosphate buffer with 500 mMNaCl, pH 7.4), then 200 nM of MMP-9 (1 μL) was added and reacted for 16hours at 37° C. 10 μL of activated MMP-9 was added to the proteinsample, after 60 and 120 minutes respectively, the samples werecollected. MALDI-TOF was utilized to measure the molecular weight ofEGF-MMP.

Melt Coextrusion of PCL Fibers

The melt coextrusion process began with PCL (CAPA 6800 pellets, MW=80kg/mol) and PEO. In order to match the rheology of PCL and PEO for themelt extrusion processing, two different molecular weights of PEO (DowChemical, POLYOX N80 (MW=200 kg/mol) and POLYOX N10 (MW=100 kg/mol) wereused with a ratio of 30:70 (200 kg/mol:100 kg/mol, N80:N10). The mixtureof two molecular weights of PEO was used to ensure a viscosity matchbetween PEO and PCL, critical to maintaining fiber uniformity. The twogrades of PEO were pre-mixed using a Haake Rheodrive 5000 twin screwextruder and pelletized. The viscosities of the obtained PEO blend andPCL melt matched at 180° C., which was chosen as the extrusiontemperature. PEO and PCL were completely dried at 40° C. under highvacuum for 48 h in advance of co-extrusion to prevent void volumeformation from residual moisture. PCL fiber domains embedded in a PEOmatrix were fabricated via multilayer co-extrusion at 180° C. 18vertical multipliers and 5 horizontal multipliers were utilized in thisprocess. Finally, this structure went through a 3″ exit die, and theextruded tape contained 8192 by 32 fiber domains. The extruded tape wascollected on a chill roll at room temperature with a speed of 15 rpm.

Preparation of Non-Woven PCL Fiber Mats

To remove the separating PEO domains, the composite tape was cut (10 cmlength) and placed in a water bath while stirring (12 hours). Afterprolonged immersion the majority of the PEO was removed from the PCL/PEOcomposite tape yielding PCL fiber bundles. Two strips of PCL fiberbundles were stacked in a cross-ply (90° C.) on a metal plate andcovered by an aluminum grid of mesh size (250 μm). A high pressurewaterjet, through a 0.010″ diameter nozzle, was swept across the fibersparallel to the directions of the fibers on the top and bottom of thefiber mat for 5 minutes each with a 500 psi rotator pressure. Thisprocess removed the remaining residual PEO and produced a non-wovenfiber mat. The pore size of the fiber mats was determined by aporometer. The specific surface area of the fiber mat was analyzed viamultipoint Brunner-Emmett-Teller (BET) analyzer (Micrometrics, TriStarIII) after degassing at 40° C. under nitrogen for 24 h.

Photochemistry

20 mg of aminooxy benzophenone, was dissolved in methanol (MeOH) (10 mgml⁻¹). Non-woven PCL fiber mats were cut to 1 cm×1 cm, width and length.Each sample was soaked in the aminooxy benzophenone solution for 5 minand then air-dried at room temperature. The dried samples were placed ona glass slide and irradiated using a UV source with a 320-390 nm filterfor 10 minutes and the fiber mat was flipped and irradiated again(Intensity=33.5 mW cm−2, n=3). Unreacted aminoxy benzophenone wasremoved by washing the fiber mats in MeOH overnight for a total of 3washes and then the samples were vacuum dried.

Transamination of EGF by PLP

200 mM of pyridoxal 5′-phosphate (PLP) was dissolved in PBS buffer andtitrated with 6N NaOH to pH 6.5. A 1:1 (protein:PLP, v/v) solution (thefinal concentrations of PLP and the protein were 100 mM and 5 mM,respectively) was made with a final pH of 6.5 and incubated at 37° C.for 1 hour. After incubation, the solution was filtered using 3.5K MWCOcentrifuge filter. The purification proceeded until no residual PLP wasobserved in the flow-through by UV-Vis spectroscopy at 414 nm. In orderto determine the success of the PLP reaction, the presence of the ketoneon the protein EGF-MMP (or control EGF) was probed by MALDI which wasused to investigate the molecular weight of the unmodified andketone-modified protein. In order to probe ketone formation, ketonefunctionalized protein was conjugated to aminooxy-5(6)-TAMRA dye in thepresence of 10 mM of aniline, incubating at 37° C. After 2 hours, extraTAMRA dye was filtered with 3.5 kDa cut off spin filter. Proteinconjugated with aminooxy-5(6)-TAMRA was analyzed with fast proteinliquid chromatography (FPLC) monitoring at 555 nm, which corresponds toTAMRA dye emission.

Bioconjugation of EGF Protein to Polymer Fiber Mat

Oxime chemistry was used to conjugate ketone-EGF (either control EGF orEGF-MMP) to the aminooxy amine-modified-PCL fiber mat. First, the fibermats were placed in 20 mL scintillation vials and a solution of EGF (60μg/mL) was added to the vial. 1% v/v of aniline was added as a catalystand the reaction proceeded for 2 hours at 37° C. After 2 hours, thefiber mats were removed from the EGF solution and rigorously washed withPBS buffer to completely remove free EGF and catalyst. Complete removalof excess protein was monitored by measuring the waste PBS at 280 nm viaUV-Vis spectroscopy. For all FPLC experiments, 2 column volumes ofmobile phase (50 mM phosphate buffer, 150 mM NaCl, pH 7.4) was passed ata flow rate of 0.4 mL/min.

ATR-FTIR Measurement for Affinity of EGF-MMP on the Alkoxyamine PCLFiber Mat

1 cm×1 cm of PCL fiber mat was prepared and soaked in 20 mg of aminooxybenzophenone in MeOH in the same manner described previously. Only oneside of the sample was exposed to the UV source and then washed in pureMeOH to remove unreacted aminooxy benzophenone. To confirm the specificattachment of ketone functionalized EGF-MMP to the aminooxybenzophenone-PCL fiber mat, oxime click chemistry was performedfollowing the same bioconjugation procedure previously described. Afterwashing and drying of the fiber mat, ATR-FTIR was utilized toinvestigate the presence of EGF-MMP on both sides of the PCL fiber mat.

EGF Release Kinetic by MMP-9

Control EGF or EGF-MMP conjugated fiber mats were prepared (n=3) storedin 2 mL of PBS buffer. In order to activate MMP-9 enzyme, amino-phenylmercuric acetate (APMA) (3.5 mg, 10 mmol) was dissolved in 1 ml of DMSOand then the solution was diluted with reaction buffer (50 mM Tris-HCLpH 7.4, 1 mM CaCl2, 0.05% Triton X-100) to a final concentration of 2 mMAPMA in solution. 100 μL of APMA solution was added to 900 μL ofphosphate buffer (50 mM phosphate buffer with 500 mM NaCl, pH 7.4) andthen, 200 nM of MMP-9 (1 μL) was added and reacted for 16 hours at 37°C. The protein conjugated PCL fiber mat was soaked in 1 mL of phosphatebuffer in an Eppendorf tube (n=3). 10 μL of activated MMP-9 was added toeach sample, after 10 minutes the buffer was collected and 1 mL of freshphosphate buffer with 10 μL of activated MMP-9 solution was added tofiber mats. This process was repeated at time points of 10, 30, 60 and120 minutes.

The released EGF samples from the fiber mat conjugated with eithercontrol EGF or EGF-MMP was evaluated via indirect ELISA. An EGF standardcurve was generated using concentrations from 100 to 1 ng/mL ofrecombinant human EGF to determine the concentration of the releasedprotein and released samples were plated at 1× or 10× dilutions. NuncMaxisorp 96-well plates were coated with 50 μL of EGF at 4° C.overnight.

PBS, pH 7.4) at room temperature for 2 hours. The wells were thenincubated with mouse anti-EGF IgG at 5 μg/mL in 100 μL blocking bufferfor 2 hours at room temperature. The wells were then incubated with 100μL of 100 ng/mL of horseradish peroxidase labeled goat anti-mouse IgG inblocking buffer for 2 hours at room temperature. The wells were washedbetween each incubation step using 4×300 μL of 0.1% w/v Tween-20 in PBS,pH 7.4. The wells were developed using 100 μL of 1-step TMB substrate at4° C. for 10 minutes. The reaction was stopped with 50 μL of 2 M H2SO4and the absorbance was read at 450 and 540 nm in triplicate for eachsample. The absorbance at 450 nm was subtracted from the absorbance at540 nm and the EGF concentration in the released sample was determinedvia comparison to the standard curve and correction via the dilutionfactor if necessary.

Cell Viability Test

HaCaT and A431 cells were grown and maintained in Dulbecco's ModifiedEagle Medium (DMEM) supplemented with 10% newborn calf serum (NCS)(Omega Scientific), 1 mM sodium pyruvate, 1% of penicillin/streptomycin,and 2 mM GlutaMax. Cells were grown at 37° C. in a humidified 5% CO₂ and95% air atmosphere. Cells used for in-vitro experiments were used atpassage numbers less than 15. A431 and HaCaT cells were plated in96-well plates in triplicate (104 cells/well) in 100 μL of completeDMEM. After 18 hours, the media was replaced with serum free media andincubated for 6 hours. Following serum starvation, EGF prepared inserum-free DMEM (100 μL/well) was added at the indicated concentrationsto the cells and allowed to incubate for 72 hours at 37° C. Cells werethen assayed for viability using the MTT assay. MTT (5 mg/mL in DPBS)was combined with complete DMEM (85:15 DMEM: MTT, 25 μL/well) and addedto each individual well and incubated at 37° C. for approximately 2hours (the assay was stopped when significant accumulation of purpleformazan crystals was visibly observed in control wells). Media wascarefully aspirated and DMSO was added (200 μL/well) to dissolve thepurple MTT-formazan crystals. Absorbance of the dissolved formazan wasquantified at 570 nm using a UV-Vis plate reader and cell viability wasdetermined as a fraction of absorbance relative to untreated controlwells. The average values are presented with standard deviation.

Scratch Cell Migration Test

Fiber mats were sterilized with 70% ethanol and dried, then a fiber matwas fixed on the side wall of each well in a 12 well culture plate withnail polish. HaCaT cells grown in DMEM supplemented with 10% FBS and 1%penicillin/streptomycin were seeded onto each well of 12 well tissueculture plate (1×10⁶ cells per well). Cell confluence reached around 80%as a monolayer after 48 hours. The monolayer was gently and slowlyscratched with a sterilized 10 μL pipette tip across the center of thewell in a straight line in one direction. The resulting scratch wasimaged via optical microscopy. Each well was washed twice with PBS toremove detached cells and the wells were replenished with serum freemedium. 10 μL of activated MMP-9 was added to each well. After 24 hours,cells were imaged again and ImageJ was utilized to measure scratchdistance.

From the above description of the invention, those skilled in the artwill perceive improvements, changes and modifications. Suchimprovements, changes and modifications within the skill of the art areintended to be covered by the appended claims. All references,publications, and patents cited in the present application are hereinincorporated by reference in their entirety.

Having described the invention, I claim:
 1. A polymer nanofiber scaffoldcomprising: a plurality of melt extruded polymer nanofibers, thenanofibers each having a rectangular cross-section defined in part by anencapsulating polymer material that is separated from the nanofibers,the nanofibers including a plurality of click-reactive functional groupsextending from portions of outer surfaces of the nanofibers, thefunctional groups being chemically bound to the nanofibers withoutdegrading polymers chains of the nanofibers.
 2. The scaffold of claim 1,wherein concentration of functional groups extending from at least oneportion is at least about 0.1 nmol/cm².
 3. The scaffold of claim 1,wherein the functional groups are spatially arranged on the nanofiberssuch that a first portion of the nanofibers has first concentration offunctional groups and second portion of the nanofibers has a secondconcentration of functional groups different than the concentration offirst portion.
 4. The scaffold of claim 1, wherein the functional groupsare spatially arranged on the nanofibers such that different portions ofthe nanofibers have different concentrations of the functional groups.5. The scaffold of claim 1, wherein the functional groups are spatiallyarranged on the nanofibers in a concentration gradient.
 6. The scaffoldof claim 1, the plurality of click-reactive function groups including afirst click reactive functional groups and second click reactivefunctional groups different than the first click reactive functionalgroups.
 7. The scaffold of claim 1, wherein the functional groups arechemically bound to the nanofibers with diarylhydroxymethylene linkagesthat are formed by reaction of a click-reactive functional groupsubstituted diarylketone with the polymer chains of the nanofibers. 8.The scaffold of claim 4, the click-reactive functional groups comprisingat least one of an amine, sulfate, thiol, hydroxyl, azide, alkyne,alkene, carboxyl groups, aldehyde groups, sulfone groups, vinylsulfonegroups, isocyanate groups, acid anhydride groups, epoxide groups,aziridine groups, episulfide groups, —CO₂N(COCH₂)₂, —CO₂N(COCH₂)₂,—CO₂H, —CHO, —CHOCH₂, —N═C═O, —SO₂CH═CH₂, —N(COCH)₂, —S—S—(C₅H₄N) andgroups of the following structures,

wherein R₁ is hydrogen or C₁ to C₄ alkyl.
 9. The scaffold of claim 1,wherein the nanofibers are formed of a polycaprolactone.
 10. Thescaffold of claim 1, wherein the encapsulating polymer material is awater soluble polymer.
 11. A method of forming an agent functionalizedpolymer nanofiber scaffold, the method comprising: providing a pluralityof melt extruded polymer nanofibers, the nanofibers each having arectangular cross-section defined in part by an encapsulating polymermaterial that is separated from the nanofibers, chemically bonding aplurality of click-reactive functional groups of a specific binding pairto the fibers without degrading polymer chains of the nanofibers, theclick-reactive functional groups extending from portions of outersurfaces of the nanofibers; and appending at least one agent to thenanofibers by reacting an agent conjugated to a complementaryclick-reactive group of the specific binding pair with theclick-reactive functional groups of the nanofibers.
 12. The method ofclaim 11, wherein the agent comprises at least one a bioactive agent,diagnostic agent, therapeutic agent, catalyst, charged molecule,peptide, polypeptide, nucleic acid, polynucleotide, small molecule,nanoparticle, antibody, carbohydrate, or vector.
 13. The method of claim11, the nanofibers are formed by: coextruding a first polymer materialwith a second polymer material to form a coextruded polymer film havingdiscrete overlapping layers of polymeric material; multiplying theoverlapping layers to form a multilayered composite film; and separatingthe first polymer material from the second polymer material to form theplurality of nanofibers having the rectangular cross-section.
 14. Themethod of claim 13, wherein separating the polymer materials includessubjecting the multilayered composite film to a high pressure waterstream.
 15. The method of claim 13, wherein separating the polymermaterials includes subjecting the multilayered composite film to a highpressure air stream.
 16. The method of claim 13, dissolving the secondpolymer material.
 17. The method of claim 11, wherein concentration offunctional click-reactive function groups extend from at least oneportion is at least about 0.1 nmol/cm².
 18. The method of claim 11,wherein the functional groups are spatially arranged on the nanofiberssuch that a first portion of the nanofibers has first concentration offunctional groups and second a portion of the nanofibers has a secondconcentration of functional groups different than the concentration offirst portion.
 19. The method of claim 11, wherein the functional groupsare spatially arranged on the nanofibers such that different portions ofthe nanofibers have different concentrations of the functional groups.20. The method of claim 11, wherein the functional groups are spatiallyarranged on the nanofibers in a concentration gradient.